Novel Electrophilic and Photoaffinity Covalent Probes for Mapping the Cannabinoid 1 Receptor Allosteric Site(s)

This is an open access article published under an ACS AuthorChoice License, which permits

copying and redistribution of the article or any adaptations for non-commercial purposes.

Article

pubs.acs.org/jmc

Novel Electrophilic and Photoaﬃnity Covalent Probes for Mapping

the Cannabinoid 1 Receptor Allosteric Site(s)

Pushkar M. Kulkarni,^‡Abhijit R. Kulkarni,^‡Anisha Korde,^‡,∥Ritesh B. Tichkule,^∥,#Robert B. Laprairie,^$

Eileen M. Denovan-Wright,^$Han Zhou,^∥,#David R. Janero,^‡,∥Nikolai Zvonok,^∥

Alexandros Makriyannis,^‡,∥,#Maria G. Cascio,^§Roger G. Pertwee,^§and Ganesh A. Thakur*^,‡

∥

#

^‡Department of Pharmaceutical Sciences, Bouve

́

College of Pharmacy, Center for Drug Discovery, and Department of Chemistry

and Chemical Biology, Northeastern University, Boston, Massachusetts 02115, United States

^$Department of Pharmacology, Dalhousie University, Halifax NS Canada B3H 4R2

^§School of Medical Sciences, Institute of Medical Sciences, University of Aberdeen, Foresterhill, Aberdeen, AB25 2ZD, Scotland

S

* Supporting Information

ABSTRACT: Undesirable side eﬀects associated with orthos-

teric agonists/antagonists of cannabinoid 1 receptor (CB1R), a

tractable target for treating several pathologies aﬀecting

humans, have greatly limited their translational potential.

Recent discovery of CB1R negative allosteric modulators

(NAMs) has renewed interest in CB1R by oﬀering a

potentially safer therapeutic avenue. To elucidate the CB1R

allosteric binding motif and thereby facilitate rational drug

discovery, we report the synthesis and biochemical character-

ization of ﬁrst covalent ligands designed to bind irreversibly to

the CB1R allosteric site. Either an electrophilic or a

photoactivatable group was introduced at key positions of two classical CB1R NAMs: Org27569 (1) and PSNCBAM-1 (2).

Among these, 20 (GAT100) emerged as the most potent NAM in functional assays, did not exhibit inverse agonism, and

behaved as a robust positive allosteric modulator of binding of orthosteric agonist CP55,940. This novel covalent probe can serve

as a useful tool for characterizing CB1R allosteric ligand-binding motifs.

agonists have been associated with adverse events including

mood alteration (euphoria, anxiety, panic), acute psychoses,

and impaired cognition and motor performance, which limit

their clinical utility.²¹Several CB1R-selective antagonists/

inverse agonists have also emerged as potential drugs for

cardiometabolic diseases and nicotine- and alcohol-use

disorders. Reminiscent of CB1R orthosteric agonists, however,

therapeutic application of CB1R orthosteric antagonists/inverse

agonists is severely restricted by the potential for unacceptable

psychotropic side eﬀects including depression, social aversion,

and suicidal ideation.^3,21−25

As has been demonstrated for several other class-A GPCRs,

CB1R has allosteric sites spatially distinct from the orthosteric

ligand-binding pocket, and allosteric modulators with CB1R

selectivity vs CB2R have been identiﬁed.²⁶⁻²⁹Engagement of

CB1R by allosteric modulators is believed to induce a

conformational change in the receptor that may be diﬃcult

to achieve with orthosteric ligands alone and “ﬁne-tune” the

pharmacological activity of the orthosteric ligand.³⁰⁻³²Due to

their generally enhanced CB1R selectivity, reduced inter-

receptor promiscuity, and higher-resolution functional control

INTRODUCTION

■

Constituents of the endocannabinoid biosignaling system

include two principal cannabinoid G-protein-coupled receptors

(GPCRs) 1 and 2 (CB1R and CB2R, respectively), their main

endogenous cannabinergic ligands (anandamide, AEA; 2-

arachidonoylglycerol, 2-AG), and enzymes responsible for

endocannabinoid biosynthesis and inactivation.¹⁻⁴Expressed

in various peripheral tissues, CB1R is the most abundant class-

A GPCR in brain.^5,6CB1R-mediated signaling helps regulate

many important physiological functions including learning,

memory, and cognition, nociception, cardiovascular function,

reproduction, and neuronal development. Dysregulated CB1R

activity has been implicated in the pathogenesis of disease states

related to these and other physiological processes such that

small-molecule modulators of CB1R-mediated signaling are

considered to have therapeutic potential.^1,3On the other hand,

CB2R is mainly expressed in peripheral tissues, particularly

immune cells^7,8as well as CNS microglia⁹and has been

pursued for treating pain and inﬂammation.¹⁰⁻¹⁷In the past

two decades, structurally diverse, potent, and selective CB1R

orthosteric agonists have been identiﬁed with (pre)clinical

eﬃcacy in treating nausea, emesis, and multiple sclerosis and

managing glaucoma, pain, and inﬂammatory disorders.¹⁸⁻²⁰

Their salutary eﬀects notwithstanding, CB1R orthosteric

Received: August 21, 2015

A

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

Figure 1. Representative CB1R allosteric modulators reported in the literature.

of receptor information transmission, CB1R allosteric modu-

lators are anticipated to oﬀer several therapeutic advantages

over orthosteric ligands.

date, very limited in vivo studies with 1 and 2 have been

reported where these NAMs have shown moderate eﬃ-

cacy.^27,38−40Another major limitation associated with these

two compounds is that they exhibit CB1R inverse agonism in

addition to having NAM activity.^26,41,42For establishing

therapeutic utility of CB1 NAMs, there is a need for developing

potent and eﬃcacious CB1R NAMs that lack inverse agonism

so as to avoid related side eﬀects.

Exemplars of well-studied, structurally distinct CB1R-

selective allosteric ligands are shown in Figure 1. These include

5-chloro-3-ethyl-N-(4-(piperidin-1-yl)phenethyl)-1H-indole-2-

carboxamide (1, Org27569)²⁶and 1-(4-chlorophenyl)-3-(3-(6-

(pyrrolidin-1-yl)pyridine-2-yl)phenyl)urea (2, PSNCBAM-1),²⁷

two CB1R allosteric modulators that emerged from initial

structure−activity relationship (SAR) studies on high-through-

put screening (HTS) leads. Although 1 and 2 exhibit several

characteristic properties of allosteric modulators, they elicit

markedly divergent eﬀects on the aﬃnity and eﬃcacy of the

standard cannabinoid-receptor orthosteric ligand 2-

[(1R,2R,5R)-5-hydroxy-2-(3-hydroxypropyl)cyclohexyl]-5-(2-

methyloctan-2-yl)phenol (CP55,940). Compounds 1 and 2

behave as both a positive allosteric modulator (PAM) of

CP55,940 binding aﬃnity and a negative allosteric modulator

(NAM) of CP55,940 signaling eﬃcacy and potency. Addition-

ally, endogenous CB1R allosteric modulators have been

identiﬁed and characterized. The nonclassical eicosanoid

(5S,6R,7E,9E,11Z,13E,15S)-5,6,15-trihydroxyicosa-7,9,11,13-

tetraenoic acid (3, lipoxin A4), whose traditional biological

target is the formyl peptide receptor FPR1, was also shown to

function as CB1R PAM of orthosteric ligand binding and

adenylyl cyclase activity.²⁸The endogenous steroid 1-

((3S,8S,9S,10R,13S,14S,17R)-3-hydroxy-10,12,13-trimethyl-

2,3,4,7,8,9,10,11,12,13,14,15,16,17-tetradecahydro-1H-

cyclopenta[a]phenanthren-17-yl)ethan-1-one (4, pregneno-

lone) acts as a CB1R NAM functionally (CB1R-mediated

ERK1/2 phosphorylation) without any eﬀect on orthosteric

agonist binding aﬃnity.²⁹The dopamine transport inhibitor

RTI-371³³and the PPAR-α agonist fenoﬁbrate have also been

suggested to act at a CB1R allosteric site.³⁴Very recently, we

have shown that cannabidiol, the nonpsychoactive constituent

of Cannabis sativa, exhibits negative allosteric modulation at

CB1R.³⁵Additionally, ligands displaying positive allosteric

modulation of orthosteric ligand’s binding and function have

been reported recently.^36,37These collective ﬁndings substan-

tiate the existence and functional signiﬁcance of allosteric sites

on CB1R whose pharmacological modulation of endogenous/

orthosteric ligand activity could be exploited for therapeutic

ends.

To inform rational drug design aimed at therapeutic CB1R

allosteric modulation, it is critical to expand our currently

limited knowledge of the structural properties and functional

inﬂuence of the receptor’s allosteric ligand-binding site(s).

Although candidate atomic-level interactions involved in GPCR

ligand recognition and functionally productive engagement can

be extrapolated from ligand−receptor cocrystals, a CB1R

crystal structure has remained elusive, and its inherently static

nature precludes direct observation of structure−function

correlates of CB1R (allosteric) ligands. Although homology

modeling and mutation studies have allowed some character-

ization of the properties of CB1R’s allosteric ligand-binding

domain, these approaches per se cannot aﬀord direct

experimental observation of the molecular nature of ligand−

CB1R interaction and its consequences for cell signaling, since

even conservative, single amino-acid mutations may alter

inadvertently receptor conformation and function.⁴¹⁻⁴⁴We

have incorporated an alternative approach for interrogating

directly the structure−function correlates of ligand binding to

druggable protein targets (enzymes, GPCRs) in their functional

state and under physiological conditions.⁴⁵⁻⁴⁹Globally, this

experimental paradigm, termed ligand-assisted protein structure

(LAPS), integrates information from point mutations, molec-

ular modeling, and peptide-level tandem mass spectrometry

studies on ligand−receptor complexes to identify amino acid

residues within (or in the immediate vicinity of) the ligand-

binding domain critical to ligand engagement and activity.⁴⁹

Pharmacologically active ligands of diverse chemical classes

purpose-designed to carry reactive groups as high-aﬃnity, site-

directed covalent probes are key elements foundational to the

LAPS experimental paradigm. Various reactive groups, both

electrophilic (e.g., isothiocyanate, benzophenone, etc.) and

photoactivatable (triﬂuoromethyl diazirine, aliphatic/aromatic

azides, etc.) type, can be incorporated at key positions into a

noncovalent parent ligand to render the parent ligand capable

of reacting in a chemically deﬁned manner with a distinct

amino acid specie.⁵⁰⁻⁵²For this purpose, and in recognition of

the importance of cysteine residues to protein structure and

Although it has been a decade since the ﬁrst CB1 NAM (1)

was reported, no new CB1 NAM with improved potency/

eﬃcacy has been identiﬁed that has been studied in vivo. To

B

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

a

Scheme 1. Synthesis of Piperidinyl Phenethylamine 8

a

Reagents and conditions: (a) piperidine, K₂CO₃, anhyd NMP, 135 °C, 12 h; (b) CH₃NO₂, NH₄OAc, reﬂux, 2 h; (c) (i) NaBH₄, MeOH, 5 °C to rt,

2 h; (ii) NiCl₂·6H₂O, NaBH₄, THF:MeOH (95:5), 0 °C, 3 h.

a

Scheme 2. Synthesis of Substituted Indole-2-carboxylates 15a, 15b, and 15c

a

Reagent and conditions: (a) NaNO₂, HCl, 0 °C, 1 h; (b) NaOEt, EtOH, 12 h, reﬂux; (c) CH₃COONa, EtOH, 0 °C, 3 h; (d) 20% H₂SO₄, EtOH,

reﬂux, 24 h.

function, we have successfully exploited the spontaneous,

preferential reactivity at physiological pH between isothiocya-

nate (NCS)-functionalized electrophilic ligands and target-

protein cysteine nucleophiles.^46,53−56The isothiocyanate

functionality exhibits pronounced reactivity toward amine

nucleophiles and sulfhydryl groups of cysteine with poor ability

to react with other nucleophiles such as alcohol or water.

The successful design and utilization of covalent aﬃnity

probes in LAPS and other experimental applications to help

characterize experimentally the orthosteric ligand-binding

domains of CB1R and other cannabinoid-system protein

therapeutic targets^{60−62,47,57,58}prompted the current work

aimed at generating a focused library of electrophilic and

photoaﬃnity probes carrying covalently reacting groups and

targeted to the CB1R allosteric site(s). The design approach

was based on the rational derivatization of two well-studied

CB1R allosteric ligands, 1 and 2 (Figure 1), guided by the

existing SAR data. Our results document the successful

extension of the application of orthosteric CB1R covalently

reactive probes to the receptor’s allosteric site(s) and constitute

the ﬁrst identiﬁcation and functional proﬁling of a novel,

covalent, allosteric CB1R aﬃnity probe.

(isothiocyanate) or photoaﬃnity (azide or benzophenone)

warheads placed at the terminal carbon of the C3 side chain

and at the C5 position on parent molecule 1, and at the C-4

positon of 2 were synthesized, generating the novel analogues

19, 20, 25, 26, 33, 34, and 36 containing covalently reacting

groups (Schemes 1−5).

The novel indole-2-carboxamide analogues (19, 20, 25, and

26) of 1 were constructed as shown in Schemes 3 and 4. The

C5-substituted indole rings were synthesized with an eﬃcient

method that utilizes Fisher cyclization on a mixture of azo and

hydrazone of the corresponding diazonium salts (Scheme 2).

The ﬁnal carboxamide derivatives were synthesized using

carbodiimide based amidation of the substituted indole-2-

carboxylic acids with piperidinyl phenethylamine synthesized as

per Scheme 1.

N-Arylation of commercially available 4-ﬂuorobenzaldehyde

(5) with piperidine gave 4-piperidinylbenzaldehyde (6).⁶⁵To

access the substituted nitrostyrene 7, we employed the Henry

reaction on 6 in the presence of ammonium acetate in

nitromethane as a solvent. Direct conversion of 7 to the desired

amine 8 using LiAlH₄, according to a previously published

protocol, required 48 h, and the product was isolated in low

yield.⁶⁶Alternatively, a route involving ﬁrst the reduction of the

double bond on 7 with NaBH₄followed by reduction of nitro

group with in situ-generated nickel borohydride gave 8 in high

yield (Scheme 1).

CHEMISTRY

■

From precedent structure−activity relationship studies con-

ducted by us⁴⁰and others⁵⁹⁻⁶⁴on the two CB1R allosteric

modulators 1 and 2, we identiﬁed two sites on the molecule,

the C3 and C5 positions of 1, and the C-4 position of 2,

important to the overall allosteric activity of these compounds.

A focused library of seven analogues with electrophilic

Scheme 2 describes the synthesis of the key substituted-

indole esters. The alkylation of ethyl acetoacetate using

(substituted) alkyl halides (11) in the presence of sodium

ethoxide gave β-ketoesters (12) as the ﬁrst step. Condensation

C

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

a

Scheme 3. Synthesis of 3-Ethylindole-2-carboxamides 19 and 20

a

Reagents and conditions: (a) dioxane:H₂O (10:1), KOH, reﬂux, 2 h, acidic workup; (b) EDCI, HOBt, DIPEA, NMP, rt, overnight; (c) NiCl₂·

6H₂O, NaBH₄, THF:CH₃OH (13:1), −5 °C, 1 h; (d) t-BuONO, TMSN₃, THF, rt, 3 h, (e) di(2-pyridyl)thionocarbonate, CH₂Cl₂, rt, 15 min.

a

Scheme 4. Synthesis of 5-Chloroindole-2-carboxamides 25 and 26

a

Reagents and conditions: (a) (i) ethanolamine, EtOH, reﬂux, 14 h; (ii) KOH, dioxane:H₂O (4:1), reﬂux, overnight; (b) Boc-anhydride, THF, aq

NaHCO₃soln, 0 °C (3 h) to rt (24 h); (c) EDCI, HOBT, DIPEA, NMP, rt, overnight; (d) TFA:CH₂Cl₂(1:10), rt, 3 h; (e) K₂CO₃, CuSO₄,

CH₃OH:H₂O (20:1), TfN₃in CH₂Cl₂, rt, 18 h; (f) di(2-pyridyl)thionocarbonate, CH₂Cl₂, rt, 15 min.

of 12 with freshly prepared diazonium salts 10 (obtained from

substituted anilines 9) in the presence of sodium acetate gave a

mixture of azos (13) and hydrazones (14) as per the Japp−

Klingeman reaction. These intermediates (13, 14) were isolated

as a mixture by passing the reaction crude product over a small

silica gel column, and in the case of 12c the reaction yielded

only the azo compound 13c.⁶⁷This was followed by Fisher

cyclization in 20% ethanolic sulfuric acid to give the chloro

indole ester (15a), the nitro indole ester (15b), and the

phthalimido indole ester (15c) in 57−71% yields.

The azido (19) and isothiocyanate (20) analogues of 1 were

constructed as depicted in Scheme 3. Base-catalyzed hydrolysis

of the nitro indole ester (15b) gave acid (16b) in high yield.

Coupling of 16b with 8 yielded the nitro indole-2-carboxamide

(17). Similarly, 15a was used to synthesize 1. The nitro

functionality of 17 was eﬃciently reduced to the amino indole-

2-carboxamide (18) using in situ-generated nickel borohydride.

Treatment of 18 with a mixture of tert-butyl nitrite and azido

trimethylsilane yielded 5-azido-3-ethyl-N-(4-(piperidin-1-yl)-

phenethyl)-1H-indole-2-carboxamide (19). Compound 18

was also served as a precursor to synthesize the corresponding

D

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

a

Scheme 5. Synthesis of Diarylureas 33, 34, and 36

a

Reagent and conditions: (a) neat, rt, 1 h; (b) Ba(OH)₂, Pd[P(Ph)₃]₄, DME:H₂O (5:2), 150 °C, M.W., 15 min; (c) Raney-nickel, H₂, MeOH, rt, 3

h; (d) triphosgene, Et₃N, toluene, 70 °C, 3 h; (e) Et₃N, DCM, 0 °C, 6 h; (f) (i) TPP, reﬂux, 4 h, benzene; (ii) CS₂, 40 °C, 12 h; (g) triphosgene,

Et₃N, toluene, 70 °C, 3 h; (h) THF, 0 °C to rt, 6 h.

isothiocyanate analogue (20) in the presence of di-2-pyridyl

thionocarbonate (DPT) at room temperature.

RESULTS AND DISCUSSION

■

Following the discovery of 1, substantial SAR studies around it

revealed that the indole-2-carboxamide scaﬀold is a promising

template through which CB1R allosteric modulators with

improved aﬃnity, eﬃcacy, potency, and pharmacokinetics

could be generated. These studies identiﬁed several key

pharmacophoric features within this structural class that

inﬂuence their binding and functional properties.^59,61−63For

example, the indole ring is more important for ligand aﬃnity

with the allosteric site than for its ability to modulate ligand

binding at the orthosteric site (allosteric cooperativity). Alkyl

chain length at the C3 position and the substitutions on the C5

position of the indole ring signiﬁcantly impact allosteric aﬃnity

as well as cooperativity at CB1R.^62,63Replacing the amide

linkage with ester functionality or modulating ethylene linker

length between the amide bond and the phenyl ring drastically

reduces the allosteric cooperativity toward binding.^60,62,64

Replacing the piperidinyl group with a dimethylamino group

signiﬁcantly increases the allosteric cooperativity, and groups

such as methyl, methylamino, nitro, and chloro but not ﬂuoro,

pyrrolidinyl, or 4-methylpiperazinyl were somewhat toler-

ated.^62,64Along similar lines, SAR reported around 2 from

us⁴⁰and others⁶¹has enabled identiﬁcation of critical positions

(especially the C-4 position) on this molecule that aﬀects

CB1R orthosteric ligand binding and downstream signaling.

The commonality of the key phamacophoric features of 1 and

2, their SAR trends, and their unique and paradoxical

pharmacological proﬁle at CB1R strongly suggest that both

may be acting through the same allosteric site on CB1R.

Phthalimido indole ester (15c) was treated with ethanol-

amine to give a lactam intermediate which was hydrolyzed

using KOH to give the acid (21) (Scheme 4). The amino group

in 21 was protected using Boc-anhydride to provide 22 which

upon coupling with 8 in the presence of EDCI aﬀorded 23.

TFA-mediated deprotection of the NH-Boc group on 23 led to

the amino analogue (24). The azido analogue (25) was then

obtained by treating 24 with in situ-generated triﬂuorometha-

nesulfonyl azide. Direct conversion of 24 into the correspond-

ing isothiocyanate analogue (26) was carried out at room

temperature using DPT.

Analogues of 2 (33, 34, and 36) were synthesized as shown

in Scheme 5. Treatment of 2,6-dibromopyridine (27) with neat

pyrrolidine (28; excess) gave pyrrolidinyl bromopyridine (30)

in quantitative yield. Microwave-accelerated Suzuki coupling of

29 with m-nitrophenylboronic acid in the presence of catalytic

Pd(PPh₃)₄aﬀorded intermediate 30 which was further reduced

to amine (31) in the presence of Raney-nickel under a

hydrogen atmosphere. The isocyanate intermediate was

synthesized in situ by treating 31 with triphosgene in the

presence of triethylamine and was then reacted with 4-

azidoaniline (32) in the presence of triethylamine to aﬀord

the desired product 33. It was then converted to the

isothiocyanate analogue (34) by treating 33 with triphenyl-

phosphine followed by exposure to CS₂(Staudinger/Aza-Wittig

reaction).⁶⁸Commercially available benzophenone aniline (35)

upon treatment with triphosgene in the presence of triethyl-

amine gave corresponding isocyanate which was further reacted

with 31 to yield the desired benzophenone-containing probe

(36).

E

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

FUNCTIONAL CHARACTERIZATION

■

A focused library of analogues bearing reactive warheads placed

at the terminal carbon of the C3 side chain and at C5 positions

on parent molecule 1 (analogues 19, 20, 25, and 26) and at the

C4 positon of 2 (analogues 33, 34, and 36) were biochemically

evaluated in a series of assays. As these analogues were expected

to potentially exhibit the “aﬃnity vs eﬃcacy paradox” similar to

the parent compounds,^26,41and as a CB1 NAM having

functional potency but no eﬀect on orthosteric ligand binding

has been identiﬁed,²⁹we chose to ﬁrst characterize these newly

synthesized analogues in two key functional assays. We

characterized both the parent compounds and their analogues

in CHO-K1 cells stably expressing human CB1R (hCB1R) by

using the PathHunter β-arrestin and HitHunter cAMP cell-

based functional assays. The PathHunter technology indexes

the ability of a test agent to aﬀect the recruitment and binding

of the pleiotropic scaﬀold protein β-arrestin following kinase

phosphorylation of agonist-bound CB1R, a process that

uncouples the phosphorylated CB1R from its cognate G

protein (routinely, G_i), subsequently targeting the receptor for

internalization and enabling the recruitment of signal trans-

ducers by the internalizing CB1R−β-arrestin complex. The

HitHunter assay indexes the ability of a test agent to modulate

forskolin-stimulated cellular adenylyl cyclase activity (i.e.,

cellular cAMP formation). Notably, β-arrestin-mediated signal-

ing is independent of both G proteins and classical second

messengers, whereas the HitHunter cAMP assay reﬂects signal

transduction dependent upon G-proteins and cAMP as second

messenger.^41,69,70

The parent compounds as well as their covalent analogues

inhibited cellular CB1R-dependent β-arrestin recruitment and

cAMP accumulation with nanomolar potencies (Figures 2 and

3 and Tables 1 and 2). The negative allosteric modulatory

activity of both 1 and 2 in the β-arrestin and cAMP assays is

congruent with previous observations (Table 1 and 2).⁴¹

Covalent analogues of 1 exhibited a 14- to 83-fold greater

potency, and slightly greater eﬃcacy, in inhibiting β-arrestin

recruitment as compared to their eﬀect on cellular cAMP

accumulation (Figure 2, Table 1). Similarly, covalent analogues

of 2 also exhibited greater potency in the β-arrestin vs cAMP

assay, but the magnitude of the diﬀerence (3- to 19-fold) was

not as great as that displayed by the covalent analogues of 1

(Figure 3, Table 2). Among these probes, 20 was the most

potent inhibitor of β-arrestin recruitment (EC₅₀= 2 nM) and

exhibited appreciable activity in the cAMP assay (EC₅₀= 174

nM) (Table 1). Compound 20 was more potent and eﬃcacious

than the parent compound, 1, in both β-arrestin and cAMP

assays (Figure 2, Table 1) and exhibited the highest functional

selectivity (83-fold) for β-arrestin vs cAMP. When the azide

group was attached to the terminal carbon of the ethyl chain at

the C3 position (25), the ability of the analogue to inhibit

forskolin-stimulated cAMP accumulation (EC₅₀= 1120 nM), as

well as its ability to inhibit β-arrestin recruitment (EC₅₀= 64

nM), was signiﬁcantly compromised. Interestingly, placing the

isothiocyanate group at the terminal carbon of the alkyl chain at

the C3 position (26) abrogated activity in the cAMP assay

(EC₅₀> 10 000 nM) and reduced activity in the β-arrestin assay

by 2 orders of magnitude (EC₅₀= 209 nM). Compound 19

displayed activity similar to that of 1 in the cAMP assay but had

modest activity in the β-arrestin assay (Table 1). None of the

covalent analogues of 2 displayed a better activity proﬁle than

the parent in either the β-arrestin or the cAMP assay (Table 2).

Figure 2. Antagonism of CP55,940-dependent β-arrestin recruitment

(A) and cAMP inhibition (B) by CB1R analogues of 1 in vitro. (A)

CHO-K1 PathHunter hCB1R cells were pretreated with indicated test

compounds (0−10 μM) for 30 min followed by treatment with

CP55,940 (EC₈₀) for 90 min. β-Arrestin recruitment was quantiﬁed

using the PathHunter assay. (B) CHO-K1 cAMP HitHunter hCB₁

cells were pretreated with allosteric modulators (0−10 μM) for 30 min

followed by treatment with CP55,940 (EC₈₀) for 30 min. cAMP

inhibition was quantiﬁed using the HitHunter assay. Data are

presented as % inhibition compared to maximal CP55,940 eﬀect

SEM from two independent replicates per assay. Derived data are

presented in Table 1.

Although the azide (33) and the isothiocyanate (34) analogues

showed reduced potencies in the cAMP assay (EC₅₀= 166 nM

and 118 nM, respectively) compared to parent compound 2

(EC₅₀= 71 nM), they were some 2- to 3-fold more potent than

1 (EC₅₀= 324 nM) and of comparable potency to the best

compound (20) in that series (EC₅₀= 174 nM). However, the

potencies of 33 and 34 in the β-arrestin assay were reduced by

1 order of magnitude as compared to parent compound 2.

Compound 36 containing the “bulkier” benzophenone

functionality at the C4 position showed no activity in the

cAMP assay and only residual activity in the β-arrestin assay, an

activity proﬁle justifying our decision not to pursue the

benzophenone analogue of 1.

To extend the functional proﬁling of what emerged from the

data presented above as our novel lead CB1R allosteric ligand,

20, we evaluated its activity in the guanosine 5′-O-(3-

[³⁵S]thio)triphosphate ([³⁵S]GTPγS) binding assay in mouse

brain membranes. This assay reﬂects the functional response of

GPCR ligands at the level of GDP/GTP exchange by the

ternary, agonist-activated GPCR−G protein complex, an event

that can modulate the activity of downstream eﬀector proteins.

The assay is considered reﬂective of the degree of G protein

activation following GPCR agonist engagement, an event more

F

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

progressively decreasing the E_maxin a concentration-dependent

manner (Figure 4) with more eﬃcacy compared to 1. When

administered alone, 20 was “silent” and did not display CB1R

agonism or inverse agonism in the [³⁵S]GTPγS binding assay

performed with mouse brain membranes (Figure 5). Also, 20

did not exhibit any signs of CB1R agonism or inverse agonism

in hCB1 CHO cell membranes up to 1 μM. A statistically

signiﬁcant, but much reduced, inverse agonism compared to 1

was observed only at suprapharmacological concentration (10

μM; data not shown). This compound has been extensively

studied for its CB1R NAM as well as inverse-agonist activity in

CB1R-mediated downstream signaling pathways and in diﬀer-

ent cell lines, where it consistently showed lack of inverse

agonism. The data are beyond the scope of this paper and will

be published elsewhere.

The activity proﬁle of 20 in the [³⁵S]GTPγS binding assay is

in marked contrast to that reported for parent compounds 1

and 2, which elicit CB1R inverse-agonist activity in addition to

acting as CB1R NAM.^26,41,42To the best of our knowledge, this

is the ﬁrst report of a potent CB1R NAM lacking inverse

agonism. This functional distinction between 20 and the

standard CB1R NAMs 1 and 2 carries signiﬁcant translational

and rational drug-design implications. As detailed elsewhere,

CB1R inverse agonism has been associated with peripheral

(e.g., gastrointestinal) and central (e.g., psychobehavioral)

adverse events in preclinical animal models of disease and in

humans.^21,23The proposition has thus been advanced that

agents capable of attenuating CB1R information transmission

with intrinsically limited, if any, functional potency to elicit

negative-eﬃcacy responses might display an enhanced beneﬁt-

to-risk proﬁle as therapeutics relative to conventional CB1R

antagonists/inverse agonists for diseases with a pathogenic

component of CB1R hyperactivity.

Structural comparison between 1 and 20 implies that

modiﬁcations of 1 at C5 can generate CB1R NAMs that retain

the aﬃnity−eﬃcacy proﬁle of 1 but are devoid of or exhibit

reduced inverse-agonist activity of conventional CB1R NAMs.

To date, very limited SAR studies have been carried out with

variations at the C5 position. This work identiﬁes C5 position

as the key site for potential modiﬁcations for generating future

CB1 NAMs lacking inverse-agonist activity. This conclusion is

supported by recently published mutational and computational

data indicating that electrostatic interactions and van der Waals

forces between the nitrogen in the piperidine ring of 1 and the

Figure 3. Antagonism of CP55,940-dependent β-arrestin recruitment

(A) and cAMP inhibition (B) and by CB₁R analogues of 2 in vitro.

Eﬀects on CP55,940-induced β-arrestin recruitment and cAMP

inhibition in the presence of various analogues of 2. (A) CHO-K1

PathHunter hCB₁cells were pretreated with allosteric modulators (0−

10 μM) for 30 min followed by treatment with CP55,940 for 90 min.

β-Arrestin recruitment was quantiﬁed using the PathHunter assay. (B)

CHO-K1 cAMP HitHunter hCB₁cells were pretreated with allosteric

modulators (0−10 μM) for 30 min followed by treatment with

CP55,940 for 30 min. cAMP inhibition was quantiﬁed using the

HitHunter assay. Data are presented as % inhibition compared to

maximal CP55,940 eﬀect SEM from two independent replicates per

assay. Derived data are presented in Table 2.

proximal to the GPCR itself in the biosignaling cascade than is

cAMP formation or G protein-independent β-arrestin signal-

ing.⁷¹We observed that 20 inhibited CP55,940-induced

[³⁵S]GTPγS binding to CB1R in mouse brain membranes by

Table 1

cAMP accumulation

β-arrestin recruitment

a

b

a

b

compound

X

Y

EC₅₀(95% CI)

E_max(%) SEM

EC₅₀(95% CI)

E_max(%) SEM

1

Cl

N₃

H

324 (294−482)

389 (332−459)

174 (121−252)

1120 (962−1280)

>10000

78.1 4.49

87.1 3.02

111 12.1

96.8 3.75

−8.84 1.18

9.05 (6.63−12.4)

28.2 (22.5−35.0)

2.09 (1.24−3.53)

64.0 (51.0−80.2)

209 (160−271)

105 2.10

105 2.07

103 1.38

105 2.70

107 3.94

19

20

25

26

NCS

Cl

H

N₃

NCS

Cl

a

b

NAM EC₅₀value (nM) in the presence of each designated test compound, determined using nonlinear regression analysis. Maximal NAM eﬀect,

determined using nonlinear regression analysis. Data are derived from Figure 2.

G

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

Table 2

a

b

NAM EC₅₀value (nM) in the presence of each designated test compound, determined using nonlinear regression analysis. Maximal NAM eﬀect,

determined using nonlinear regression analysis. Data are derived from Figure 3.

Figure 5. Activity of 20 and CP55,940 in the [³⁵S]GTPγS assay. Mean

% changes in [³⁵S]GTPγS binding to mouse brain membranes elicited

by CP55,940 or 20. The mean EC₅₀value of CP55,940 with a 95%

Figure 4. [³⁵S]GTPγS assay depicting the NAM eﬀect of increasing

conﬁdence interval is 27.3 nM (7.2−103.2 nM, n = 6), and its

concentrations of 20. Mean % increases in [³⁵S]GTPγS binding to

corresponding E_maxvalue is 87.8% (67.9 and 107.7%). Vertical lines

mouse brain membranes induced by CP55,940 in the presence of

show SEM values.

DMSO (n = 12) or 100 nM or 1 μM of 20 (n = 6) or 1 μM of 1 (n =

4). The mean E_maxvalue of CP55,940 with its 95% conﬁdence interval

in parentheses is 94.3% (85.8−102.8%) in the presence of DMSO,

71.7% (65.3−78.1%) in the presence of 100 nM of 20, 11.2% (5.6−

16.8%) in the presence of 1 μM of 20, and 32.4% (24.0−40.8%) in the

presence of 1 μM of 1. Vertical lines show SEM values.

membranes obtained from CHO cells overexpressing hCB1R.

Both 20 and 1 signiﬁcantly enhanced the binding of

[³H]CP55,940 to hCB₁CHO cell membranes and acted as

CB1 PAM of binding. As indicated by the data shown in Figure

6, 20 produced this enhancement with signiﬁcantly greater

potency (lower EC₅₀) than 1.

CB1R aspartate residue D6.58(366) in CB1R transmembrane

helix 6 is crucial for inverse agonism such that the absence of

this nitrogen abrogates the inverse-agonism action of 1.⁴²In

our hands, even with the presence of the piperidine nitrogen,

20 did not evidence inverse agonism in mouse brain

membranes and hCB1 CHO cells (up to 1 μM). This result

invites the notion that the increased length of the NCS group at

C5 in 20 compared to the Cl group at that position in 1 might

extend 20 in its CB1R binding pocket slightly beyond 1’s

original docking position, potentially obviating or reducing the

interaction between the C5 nitrogen on the piperidine ring of

20 and that of the CB1R D6.58(366) residue, leading to loss of

inverse agonism.

To investigate the ability of 20 to label the CB1R allosteric

site(s) covalently, we carried out time-course experiments

between 20 and hCB1R in membranes isolated from HEK293

cells overexpressing the receptor. We indexed the covalent

association between 20 and hCB1R as the extent to which a

preincubation of the isolated membranes with 20 at a

concentration of 500 nM followed by extensive membrane

washings with centrifugation inﬂuencing the subsequent level

(B_max) of speciﬁc [³H]CP55,940 binding to hCB1R in the

washed membranes. The binding of [³H]CP55,940 to hCB1R

increased in a time-dependent manner, reaching a maximum by

60 min preincubation time with isothiocyanate 20 (Figure 7).

Incubation of the CB1 receptor with 20 for 90 and 120 min

reduced speciﬁc binding of [³H]CP55,940. Presumably, the

extended incubation time (beyond 60 min) resulted in

nonspeciﬁc covalent modiﬁcation of the receptor and impaired

its ability to bind [³H]CP55,940. These data are consistent with

LIGAND BINDING STUDIES

■

To proﬁle the ligand-binding characteristics of our lead CB1R

covalent ligand, 20, we ﬁrst evaluated the eﬀect of 20 on the

speciﬁc binding of the orthosteric ligand [³H]CP55,940 to

H

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

Figure 6. Positive allosteric modulation of [³H]CP55,940 binding by

20 and 1. Eﬀects of 20, 1, and CP55,940 on [³H]CP55,940 binding to

hCB₁CHO cell membranes (n = 6). Mean values signiﬁcantly

diﬀerent from zero are indicated by the symbols * (for 20), # (for 1),

and • (for CP55,940); one symbol = P < 0.05; three symbols = P <

0.001; Student’s one sample t test). Positive values indicate

enhancement of [³H]CP55,940 binding. The mean E_maxvalues of

20, 1, and CP55,940, with their 95% conﬁdence limits shown in

brackets, are 116.5% (108.3 and 124.6%), 109.3% (99.1 and 119.5%),

and −90.1% (−81.6 and −98.6%), respectively. The corresponding

EC₅₀values, again with 95% conﬁdence limits shown in brackets, are

19.6 nM (10.4 and 36.9 nM), 83.0 nM (44.1 and 156.2 nM), and 4.9

nM (2.6 and 9.0 nM).

Figure 8. Comparative covalent labeling of 20 (A) and 1 (B) with

hCB1R. Eﬀect of (A) 20 or (B) 1 on the saturation binding of [³H]

CP55,940 to HEK293-hCB1R cell membranes. Membranes were

incubated with allosteric ligands (500 nM) at 30 °C for 60 min and

extensively washed, followed by saturation binding of [³H]CP55,940

at concentrations ranging from 0 to 25 nM. Unbound [³H]CP55,940

was removed by washing−ﬁltration, and the membrane-bound

radioactivity was quantiﬁed. 1 did not label the receptor covalently

and hence did not aﬀect CP55,940 binding after wash, whereas 20

labeled the receptor covalently and caused ∼2.25 fold increase in

CP55,940 binding, as indexed by the respective B_maxvalues. Data

shown represent the mean SEM of three independent experiments

performed in duplicate.

that preincubation with 20 increased maximal hCB1R speciﬁc

binding of [³H]CP55,940 by ∼2.25-fold, whereas 1 had no

eﬀect. The combined data in Figures 7 and 8 indicate that 20

covalently labels hCB1R by virtue of its C5 isothiocyanate

group.

Figure 7. Time-course studies showing the eﬀect of 20 preincubation

on CP55,940 binding. HEK293-hCB1R membranes incubated with

500 nM of 20 for 0 (control), 30, 60, 90, and 120 min.

the characteristic dependency of the association between

covalent ligands/probes and target proteins upon the length

of time the protein is incubated with the probe.^{60,62,47,57,58}

The data in Figure 7 provided suﬃcient guidance to establish

experimental parameters for determining the eﬀect of 20 on

[³H]CP55,940 speciﬁc binding to hCB1R over a range of

radioligand concentrations in a saturation-binding assay. Parent

compound 1 was proﬁled in parallel as the nonderivatized

control lacking the chemically reactive isothiocyanate moiety

and, thus, incapable of covalently interacting with hCB1R. In

accord with our previous receptor-labeling studies,⁴⁵⁻⁴⁸we

pretreated the HEK293 hCB1R membranes with either test

compound (500 nM, ﬁnal concn) for 60 min, subsequently

washed the membranes extensively to quench this incubation,

and quantiﬁed any change observed for the subsequent speciﬁc

binding of [³H]CP55,940 to hCB1R (as B_max) in the washed

membranes. The saturation binding curves (Figure 8) show

CONCLUSION

■

Adverse events associated with standard CB1R orthosteric

ligands (agonists and antagonists/inverse agonists) have

prompted alternative approaches in chemical pharmacology

for leveraging the translational potential of drug-like small-

molecules that modulate CB1R-dependent signaling. Predom-

inant in this newer thinking for enhancing therapeutics

targeting of CB1R are neutral antagonists with minimal, if

any, inverse-agonist eﬀects and allosteric ligands.^23,24Due to

greater receptor selectivity and ﬁner control over downstream

signaling than standard CB1R orthosteric modulators, CB1R

allosteric modulators oﬀer opportunities for novel pharmaco-

therapies with potentially enhanced safety and eﬃcacy proﬁles.

Rational design of CB1R allosteric modulators as drugs requires

greater understanding of the receptor’s allosteric binding site(s)

and the molecular pharmacology of ligands that are engaged by

I

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

it. As part of our continued commitment to address this need,

we report our approach of derivatizing the classical CB1R

allosteric modulators 1 and 2 with a chemically reactive

electrophilic (NCS) or photoactivatable (azide and benzophe-

none) groups at the judiciously selected positions within each

molecule. Functional characterization demonstrated that these

novel analogues displayed the prototypical paradoxical

pharmacological properties which make them PAMs of

CP55,940 binding but NAM in function. With all of these

analogues and in the assay systems utilized, we observed a

consistent bias toward β-arrestin over cAMP-dependent

signaling. Among these, 20 emerged as the most potent

NAM in the cellular cAMP, G-protein-independent β-arrestin,

and G protein-dependent GTPγS functional assays, and it was

more potent than the parent compound 1. It was also more

potent than 1 as a PAM of orthosteric ligand binding. Notably,

when applied alone, 20 did not aﬀect the constitutive activity of

the receptor in the GTPγS assay, making it the ﬁrst ever

covalent, potent CB1R NAM without signiﬁcant inverse-

agonist activity, a property suggestive of a lower adverse-

event risk. Compound 20 engaged the CB1R allosteric site(s)

covalently, making this compound a unique and valuable probe

with which to help elucidate the (sub)molecular features of

ligand recognition and engagement by CB1R. Ongoing work in

this regard will incorporate 20 and newer-generation CB1R

allosteric covalent probes into our established LAPS paradigm

in conjunction with site-speciﬁc CB1R point mutations and

peptide-level LC/MS/MS for identifying experimental amino

acid residues critical to CB1R allosteric ligand binding/function

and mapping structural features and topology of the CB1R

allosteric ligand-binding pocket(s). Because preliminary data

indicate that binding of 20 to hCB1R is irreversible, the utility

of this compound as a structural probe may be extended to 20−

hCB1R cocrystallization studies aimed at mapping the location

of the CB1R allosteric site(s) and their atomic-level features.

incubated at 37 °C under 5% CO₂. Compounds were dissolved

in DMSO and diluted in OCC media. Agonist EC₈₀was

determined directly from an agonist dose−response curve (data

not shown). The CP55,940 EC₈₀was 7.5 0.15 nM (mean

SEM, n = 3 independent experiments). Media was aspirated

and replaced with 10 μL of 1:1 HBSS/HEPES:cAMP XS+Ab

reagent containing 20 μM forskolin (DiscoveRx). Five

microliters of test compound or vehicle solution was added

to each well at the appropriate concentrations and incubated

for 30 min. Five microliters of agonist was then added to each

well followed by a 30 min incubation. Twenty microliters of

cAMP XS+ED/CL lysis cocktail (DiscoveRx) was then added

followed by 60 min incubation at room temperature. Finally, 20

μL of cAMP XS+EA reagent (DiscoveRx) was added followed

by 3 h incubation at room temperature. Chemiluminescence

was measured on a standard luminescence plate reader (as

RLUs). Basal RLU was deﬁned as zero. Results were calculated

as the percentage inhibition of CP55,940 maximal eﬀect. Data

were analyzed using the four-parameter variable slope and

allosteric EC₅₀shift nonlinear regression equations in Graph-

Pad Prism 5.0 (GraphPad, San Diego, CA). The results of this

analysis are presented as E_maxSEM, and EC₅₀(nM) with 95%

CI.

Radioligand Displacement Assay. Chinese hamster

ovary (CHO) cells transfected with cDNA encoding human

cannabinoid CB₁receptors were maintained at 37 °C in

Dulbecco’s modiﬁed Eagle’s medium nutrient mixture F-12

HAM, supplemented with 1 mM ^L-glutamine, 10% fetal bovine

serum, 0.6% penicillin−streptomycin, and G418 (400 μg/mL).

All cells were exposed to 5% CO₂in their media, and were

passaged twice a week using nonenzymatic cell dissociation

solution. For membrane preparation, cells were removed from

ﬂasks by scraping, centrifuged, and then frozen as a pellet at

−20 °C until required. Before use in a radioligand binding

assay, cells were defrosted, diluted in Tris buﬀer (50 mM Tris-

HCl and 50 mM Tris-base), and homogenized with a 1 mL

hand-held homogenizer.

METHODS

■

The assays were carried out with [³H]CP55,940 and Tris

binding buﬀer (50 mM Tris-HCl, 50 mM Tris-base, 0.1% BSA,

pH 7.4), total assay volume 500 μL, using the ﬁltration

procedure described previously.^41,72Binding was initiated by

the addition of transfected human CB₁CHO cell membranes

(50 μg of protein per well). All assays were performed at 37 °C

for 60 min before termination by the addition of ice-cold Tris

binding buﬀer, followed by vacuum ﬁltration using a 24-well

sampling manifold (Brandel Cell Harvester; Brandel Inc.,

Gaithersburg, MD) and Brandel GF/B ﬁlters that had been

soaked in wash buﬀer at 4 °C for at least 24 h. Each reaction

well was washed six times with a 1.2 mL aliquot of Tris binding

buﬀer. The ﬁlters were oven-dried for 60 min and then placed

in 3 mL of scintillation ﬂuid (Ultima Gold XR, PerkinElmer,

Seer Green, Buckinghamshire, UK). Radioactivity was

quantiﬁed by liquid scintillation spectrometry. Speciﬁc binding

was deﬁned as the diﬀerence between the binding that occurred

in the presence and absence of 1 μM unlabeled CP55,940. The

concentration of [³H]CP55,940 used in our displacement

assays was 0.7 nM. The compounds under investigation were

stored as 10 mM stock solutions in DMSO, the vehicle

concentration in all assay wells being 0.1% DMSO.

PathHunter CB₁β-Arrestin Assay. Chinese hamster ovary

K1 (CHO-K1)-PathHunter hCB₁β-arrestin cells (DiscoveRx,

Fremont, CA) were seeded at 5000 cells/well in 384-well plates

24 h before use and incubated at 37 °C, 5% CO₂. Compounds

were dissolved in dimethyl sulfoxide (DMSO) and diluted in

optimized cell culture (OCC) media. Agonist EC₈₀was

determined directly from an agonist dose−response curve

(data not shown). The CP55,940 EC₈₀was 31.1 0.47 nM

(mean

SEM, n = 3 independent experiments). Five

microliters of allosteric modulator or vehicle solution was

added to each well at the appropriate concentrations and

incubated for 30 min. Five microliters of agonist was then

added to each well followed by a 90 min incubation. Fifteen

microliters of detection reagent was then added followed by

further 60 min incubation at room temperature. Chemilumi-

nescence was measured on a standard luminescence plate

reader as relative light units (RLU). Basal RLU was deﬁned as

zero. Results were calculated as the percentage inhibition of

CP55,940 maximal eﬀect. Data were analyzed using the four-

parameter variable-slope and allosteric EC₅₀shift nonlinear

regression equations in Prism 5.0 (GraphPad, San Diego, CA).

The results of this analysis are presented as E_maxSEM, and

EC₅₀(nM) with 95% CI.

[³⁵S]GTPγS Binding Assay. Mouse brain membranes (5 μg

protein), prepared as described previously,⁴¹were preincubated

for 30 min at 30 °C with adenosine deaminase (0.5 IU/mL).

The membranes were then incubated with CP55,940 or 20, or

HitHunter cAMP Assay. Chinese Hamster Ovary K1

(CHO-K1)-HitHunter hCB1R cells (DiscoveRx) were seeded

at 10 000 cells/well in 384-well plates 24 h before use and

J

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

with CP55,940 20, 1, or vehicle, for 60 min at 30 °C in assay

buﬀer (50 mM Tris-HCl, 50 mM Tris-base, 5 mM MgCl₂, 1

mM EDTA, 100 mM NaCl, 1 mM DTT, 0.1% BSA) in the

presence of 0.1 nM [³⁵S]GTPγS and 30 μM GDP, in a ﬁnal

volume of 500 μL. Binding was initiated by the addition of

[³⁵S]GTPγS. Nonspeciﬁc binding was measured in the

presence of 30 μM GTPγS. The reaction was terminated by

rapid vacuum ﬁltration (50 mM Tris-HCl, 50 mM Tris-base,

0.1% BSA) using a 24-well sampling manifold (cell harvester,

Brandel, Gaithersburg, MD) and GF/B ﬁlters (Whatman,

Maidstone, UK) that had been soaked in buﬀer (50 mM Tris-

HCl, 50 mM Tris-base, 0.1% BSA) for at least 24 h. Each

reaction tube was washed six times with a 1.2 mL aliquot of ice-

cold wash buﬀer. The ﬁlters were oven-dried for at least 60 min

and then placed in 3 mL of scintillation ﬂuid (Ultima Gold XR,

PerkinElmer, Seer Green, Buckinghamshire, UK). Radioactivity

was quantiﬁed by liquid scintillation spectrometry.

Data Analysis. Most results were calculated as percentage

changes from a basal level (zero) of [³H]CP55,940 or

[³⁵S]GTPγS binding (in the presence of vehicle). GraphPad

Prism 5.0 (GraphPad, San Diego, CA) was used to construct

sigmoidal log concentration−response curves and to calculate

values of EC₅₀, E_max, SEM, and 95% conﬁdence intervals. Some

mean values were compared using Student’s one sample t test.

P values <0.05 were considered to be signiﬁcant.

The assay was performed in 200 μL of TME−BSA buﬀer at

30 °C for 1 h with gentle agitation. Filtration and washing were

performed as described above. Nonspeciﬁc binding was

subtracted from the total bound radioactivity to obtain

[³H]CP55,940 speciﬁc binding of (as pmol/mg protein). All

assays were performed in triplicate, and data points were

represented as the mean B_maxand K_dvalues, calculated by

nonlinear regression using GraphPad Prism 5.0.

EXPERIMENTAL SECTION

■

All commercial chemicals and solvents were purchased from Sigma-

Aldrich, Inc. (St. Louis, MO), Alfa Aesar, and Combi-blocks as reagent

grade and unless otherwise speciﬁed were used without further

puriﬁcation. Biotage Initiator microwave system was used for the

synthesis of a few of the intermediates of the ﬁnal covalent probes.

Reaction progress was monitored by thin-layer chromatography

(TLC) using commercially prepared silica gel 60 F254 glass plates.

Compounds were visualized under ultraviolet (UV) light or by staining

with iodine, phosphomolybdic acid, or p-anisaldehyde reagent. Flash

column chromatography was carried out on a Biotage SP1, Biotage

Isolera, or Interchim puriﬁcation unit using prepacked columns from

Reveleris, Biotage, and Luknova. Solvents used include hexanes, ethyl

acetate, acetone, methanol, and dichloromethane. Characterization of

compounds and their purity were established by a combination of

HPLC, TLC, mass spectrometry, and NMR analyses. NMR spectra

were recorded in DMSO-d₆, chloroform-d, or methanol-d₄, on a Varian

NMR spectrometer (¹H NMR at 500 MHz and ¹³C NMR at 125

MHz). Chemical shifts were recorded in parts per million (δ) relative

to tetramethylsilane (TMS; 0.00 ppm) or solvent peaks as the internal

reference. Multiplicities are indicated as br (broadened), s (singlet), d

(doublet), t (triplet), q (quartet), quin (quintet), sept (septet), or m

(multiplet). Coupling constants (J) are reported in hertz (Hz). All test

compounds were greater than 95% pure as determined by LC/MS

analysis performed using a Agilent Technologies 1260 Inﬁnity reverse-

phase HPLC, with a dual-wavelength UV−visible detector and an

Agilent Technologies 6120 Quadrupole mass spectrometer (electro-

spray ionization).

Rat Brain and HEK293 Cell Membrane Preparations.

Rat forebrain membranes for binding assays were prepared

according to a published protocol.⁷³HEK293 cells over-

expressing hCB1R were disrupted by cavitation in a pressure

cell, and membranes were sedimented by ultracentrifugation, as

described.⁴⁶The membrane pellet was resuspended in TME

buﬀer (50 mM Tris-HCl, 5 mM MgCl₂, 1 mM EDTA, pH 7.4),

and membrane protein was quantiﬁed with a Bradford dye-

binding method (Bio-Rad Laboratories).

[³H] CP55,940 Saturation-Binding to hCB1 in the

Presence of Allosteric Ligands. Membrane preparations

either from rat brain or HEK293 cells overexpressing hCB1

receptor were resuspended in TME−BSA (TME containing

0.1% BSA), and aliquots of this suspension containing 25 μg of

proteins were added to each assay well. Membranes were

preincubated with 1 (for 1 h) or 20 (for 0, 30, 60, 90, and 120

min) at 500 nM concentration of allosteric ligand at 30 °C with

agitation. The excess ligand was removed during washes with

TME−BSA and TME buﬀers and centrifugations at 27 000g, 30

°C. Membrane proteins were quantiﬁed with a Bradford dye

binding method (Bio-Rad Laboratories). Saturation binding

assays were performed with the washed membranes and

[³H]CP55,940 radioligand at concentrations ranging between 0

and 25 nM. Nonspeciﬁc binding was evaluated in the presence

of 5 μM unlabeled CP55,940. The assay was performed at 30

°C for 1 h with gentle agitation. The resultant material was

transferred to Uniﬁlter GF/B ﬁlter plates and the unbound

ligand removed using a Packard Filtermate-96 Cell Harvester

(PerkinElmerPackard, Shelton, CT). Filter plates were washed

four times with ice-cold wash buﬀer (50 mM Tris base, 5 mM

MgCl₂containing 0.5% BSA, pH 7.4). Bound radioactivity was

quantiﬁed using the Packard top count scintillation counter.

Nonspeciﬁc binding was subtracted from the total bound

radioactivity to obtain the speciﬁc binding of [³H]CP-55,940

(represented as pmol/mg of protein). All assays were

performed in triplicate, and data points were represented as

the mean B_maxand K_dvalues, calculated by nonlinear regression

using Graphpad Prism 4.0 on a Windows platform.

5-Chloro-3-ethyl-N-[4-(piperidin-1-yl)phenethyl]-1H-indole-

2-carboxamide (1). To a mixture of 16a (500 mg, 2.24 mmol) and 8

(548 mg, 2.68 mmol) in 5 mL of anhydrous NMP under an argon

atmosphere and at room temperature were added HOBT (302 mg,

2.24 mmol), DIPEA (347 mg, 2.68 mmol), and EDCI (486 mg, 3.13

mmol), and the resulting mixture was stirred overnight. The reaction

mixture was diluted with cold water and the crude product was

extracted in ether (3×). The combined organic layer was washed with

water and brine and dried (Na₂SO₄). Volatiles were evaporated under

reduced pressure, and the crude product obtained was puriﬁed by ﬂash

column chromatography on silica gel (10−40%; EtOAc:hexanes) to

give 1 as a white solid (709 mg, 77% yield). R_f= 0.7 (EtOAc/hexanes

= 50/50). ¹H NMR (500 MHz, chloroform-d): 9.28 (s, 1H), 7.54 (d, J

= 1.5 Hz, 1H), 7.29 (d, J = 9.0 Hz, 1H), 7.20 (dd, J = 8.5 Hz, J = 2.0

Hz, 1H), 7.14 (d, J = 8.5 Hz, 2H), 6.92 (d, J = 8.5 Hz, 2H), 6.04−5.94

(m, 1H,), 3.79 (q, J = 6.5 Hz, 2H), 3.13 (t, J = 5.5 Hz, 4H), 2.89 (t, J =

6.5 Hz, 2H), 2.69 (q, J = 8.0 Hz, 2H), 1.75−1.67 (m, 4H), 1.61−1.54

(m, 2H), 1.08 (t, J = 7.5 Hz, 3H). Mass spectrum m/z 410.18 [M +

H]⁺.

4-(Piperidin-1-yl)benzaldehyde (6). To a solution of 5 (20 g,

161.1 mmol) and piperidine (16.47 g, 193 mmol) in 60 mL of dry

NMP under an argon atmosphere was added anhydrous K₂CO₃(44.5

g, 322 mmol), and the resulting solution was stirred at 135 °C for 12 h.

Reaction contents were allowed to cool to room temperature and were

diluted with ice cold water. Product was extracted in ether (3×), and

the combined organic layer was washed with water and brine and dried

over MgSO₄. Solvent was evaporated under reduced pressure, and

puriﬁcation of the crude product by ﬂash column chromatography (0−

15%; EtOAc:hexanes) aﬀorded 6 as a pale yellow solid (23.78 g, 78.2%

yield). R_f= 0.5 (EtOAc/hexanes = 20/80). ¹H NMR (500 MHz,

chloroform-d): δ 9.74 (s, 1H), 7.72 (d, J = 9.5 Hz, 2H), 6.89 (d, J = 9.0

K

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

Hz, 2H), 3.45−3.37 (m, 4H), 1.74−1.63 (m, 6H). Mass spectrum m/z

190.12 [M + H]⁺.

a period of 30 min through a dropping funnel, and the reaction

mixture was reﬂuxed for 12 h. The mixture was cooled to room

temperature and ﬁltered, and the ﬁltrate was neutralized by adding 1 N

hydrochloric acid, concentrated under reduced pressure, and

partitioned in ethyl acetate and water. The organic layer was separated,

the aqueous layer extracted with ethyl acetate (3×), and the combined

organic layer was washed with brined and dried over MgSO₄. The

product was puriﬁed by ﬂash column chromatography (0%−20%;

EtOAc:hexanes) to give 12a as a clear liquid (35.1 g, 78% yield). R_f=

0.45 (EtOAc/hexanes = 20/80). ¹H NMR (400 MHz, chloroform-d) δ

4.20 (q, J = 7.2 Hz, 2H), 3.42 (t, J = 7.2 Hz, 1H), 2.22 (s, 3H), 1.92−

1.76 (m, 2H), 1.40−1.20 (m, 5H, especially 1.28, t, J = 7.2 Hz, 3H),

0.93 (t, J = 7.2 Hz, 3H). Mass spectrum m/z 172.10 [M + H]⁺.

Ethyl 2-Acetyl-4-(1,3-dioxoisoindolin-2-yl)butanoate (12c).

To a 500 mL ﬂask containing 150 mL of anhydrous acetone were

added ethyl acetoacetate (10.0 g, 77.0 mmol) and K₂CO₃(11.68 g, 85

mmol), and the mixture was stirred for 2 h at room temperature. To

this was added 2-(3-iodopropyl)isoindoline-1,3-dione (11c; 24.21 g,

77.0 mmol) and reﬂuxed overnight. The mixture was ﬁltered through a

Buchner funnel, and the ﬁltrate was cooled to room temperature.

Volatiles were evaporated, water was added to the crude product, and

the resultant mass was acidiﬁed to pH 4. The aqueous layer was

extracted in dichloromethane (2×), and the combined organic layer

was dried over MgSO₄and triturated in hexane to obtain the desired

compound 12c as a white solid (18.31 g, 75.1% yield). ¹H NMR (500

MHz, DMSO-d₆) δ 7.87−7.82 (m, 2H), 7.74−7.69 (m, 2H), 4.19 (qd,

J = 7.0 Hz, 2.0 Hz, 2H), 3.71 (t, J = 7.0 Hz, 2H), 3.50 (t, J = 7.0 Hz,

1H), 2.25 (s, 3H), 1.96−1.81 (m, 2H), 1.78−1.63 (m, 2H), 1.26 (t, J =

7.0 Hz, 3H). Mass spectrum m/z 332.15 [M + H]⁺.

General Procedure for Synthesis of Substituted Indole-2-

carboxylates. To a solution of 2-alkylated ethyl acetoacetate 12 (2.9

mmol) in 30 mL of ethanol was added NaOAc (6.12 mmol) under an

argon atmosphere, and the resulting mixture was stirred at room

temperature for 45 min, followed by cooling to 0 °C. Aryldiazonium

salt 10 was added to the reaction along with additional NaOAc to

maintain the pH at 5, and the resulting solution was stirred for 3 h

while maintaining the temperature between 0 and 5 °C. The reaction

was quenched by adding saturated aqueous NaHCO₃solution, and the

volatiles were removed under reduced pressure. The mixture was

extracted with ethyl acetate (3×), the organic layer was washed with

water and brine and dried over Na₂SO₄. The solvent was removed

under vacuum to give the crude product as red oil which was a mixture

of corresponding azo and hydrazone (13 and 14) intermediates. This

mixture was passed through a small column of silica and dried under

high vacuum to give a thick orange-brown mass. To this mass was

added 100 mL of 20% sulfuric acid in anhydrous ethanol and was

reﬂuxed for 24 h. The reaction mixture was cooled to room

temperature and neutralized by adding aqueous NaHCO₃solution.

Crude product was extracted with ethyl acetate (3×). The combined

organic layer was washed with water and brine, dried over Na₂SO₄, and

concentrated under vacuum to yield a crude mixture which was

puriﬁed by ﬂash column chromatography on silica gel (0%−20%;

EtOAc: hexanes) to give pure desired indole esters 15.

(E)-1-[4-(2-Nitrovinyl)phenyl]piperidine (7). To a solution of 6

(20 g, 106 mmol) in 100 mL of anhydrous nitromethane was added

NH₄OAc (24.44 g, 317 mmol) under an argon atmosphere, and the

resulting mixture was reﬂuxed for 2 h. Solvent was removed under

reduced pressure, and the reaction mixture was diluted with ethyl

acetate and water (2:1). The organic layer was separated, the aqueous

layer was extracted with ethyl acetate (3×), and the combined organic

layer was washed with water and brine and dried over MgSO₄. The

solvent was evaporation under reduced pressure, and the crude

product was puriﬁed by ﬂash column chromatography (5%−20%;

EtOAc:hexanes) to give 7 as a dark orange solid (16.10 g, 65.6%

1

yield). R_f= 0.48 (EtOAc/hexanes = 20/80). H NMR (500 MHz,

chloroform-d): δ 7.95 (d, J = 13.5 Hz, 1H), 7.50 (d, J = 13.5 Hz, 1H),

7.42 (d, J = 9.0 Hz, 2H), 6.86 (d, J = 8.5 Hz, 2H), 3.40−3.36 (m, 4H),

1.74−1.63 (m, 6H). Mass spectrum m/z 233.12 [M + H]⁺.

2-(4-(Piperidin-1-yl)phenyl)ethanamine (8). To a cooled (5

°C) solution of 7 (15 g, 64.3 mmol) in 120 mL of anhydrous methanol

at 5 °C was added NaBH₄(14.66 g, 387 mmol) in small portions

under an argon atmosphere, and the reaction mixture was stirred for 2

h while allowing it to warm to room temperature. It was then

quenched with dropwise addition of 80 mL of saturated NH₄Cl. The

mixture was concentrated under reduced pressure, and the crude

product was partitioned in ethyl acetate and water. The organic layer

was separated, the aqueous layer extracted with ethyl acetate (3×), and

the combined organic layer was washed with brine and dried over

Na₂SO₄. Evaporation of volatiles under reduced pressure gave a crude

mixture which was puriﬁed by ﬂash column chromatography (10%−

40%; EtOAc:hexanes) to yield the intermediate 1-(4-(2-nitroethyl)-

phenyl)piperidine as a pale yellow oil (12.8 g, 85% yield). R_f= 0.42

1

(EtOAc/hexanes = 20/80). H NMR (500 MHz, chloroform-d): δ

7.07 (d, J = 9.0 Hz, 2H), 6.88 (d, J = 9.0 Hz, 1H), 4.55 (t, J = 7.5 Hz,

2H), 3.23 (d, J = 7.5 Hz, 2H), 3.17−3.10 (m, 4H), 1.73−1.66 (m,

4H), 1.61−1.53 (m, 2H). Mass spectrum m/z 233.12 [M + H]⁺.

To a solution of this intermediate (12.5 g, 53.40 mmol) in 100 mL

of anhydrous THF and methanol (13:1) was added NiCl₂·6H₂O

(15.22 g, 64 mmol) under an argon atmosphere, and reaction mixture

was stirred for 45 min at room temperature. It was then cooled to −5

°C, and NaBH₄(12.11 g, 320 mmol) was added in small portions. The

reaction was then gradually warmed to room temperature, stirred for 3

h, quenched with saturated aqueous solution of NH₄Cl, and

concentrated under reduced pressure. The residue was diluted with

ethyl acetate and water and ﬁltered. The organic layer was separated,

and the aqueous layer was extracted with ethyl acetate (3×). The

combined organic layer was washed with brine, dried (Na₂SO₄), and

evaporated under vacuum to yield pure amine 8 (8.8 g, 81% yield). R_f

= 0.81 (MeOH/DCM = 20/80). ¹H NMR (500 MHz, chloroform-d):

δ 7.08 (d, J = 8.0 Hz, 2H), 6.89 (d, J = 8.0 Hz, 2H), 3.17−3.08 (m,

4H), 2.92 (t, J = 7.0 Hz, 2H), 2.66 (t, J = 7.0 Hz, 2H), 1.76−1.66 (m,

4H), 1.61−1.52 (m, 2H), 1.31 (br s, 2H). Mass spectrum m/z 205.16

[M + H]⁺.

4-Chlorobenzenediazonium Chloride (10a). To a suspension

of ﬁnely powdered 9a (2.54 g, 20 mmol) in 10 mL of 24% aq

hydrochloric acid at 0 °C was added a cold aqueous solution of sodium

nitrite (1.7 g, 23 mmol), and the reaction mixture was stirred for 1 h

while maintaining the temperature between 0 and 5 °C. The resulting

pale yellow solution of diazonium salt 10a was directly used for the

next reaction.

4-Nitrobenzenediazonium Chloride (10b). The compound was

synthesized as per the procedure described for 10a using nitroaniline

9b.The reaction solution of diazonium salt 10b was directly used for

the next reaction.

Ethyl 5-Chloro-3-ethyl-1H-indole-2-carboxylate (15a). The

compound was synthesized as per the general procedure as a white

solid (64% yield). R_f= 0.35 (EtOAc/hexanes = 20/80). ¹H NMR (400

MHz, chloroform-d): δ 8.74 (br s, 1H), 7.65 (d, J = 1.2 Hz, 1H), 7.30

(d, J = 8.8 Hz, 2H), 7.25 (dd, J = 8.8 Hz, J = 1.2 Hz, 1H), 4.42 (q, J =

7.2 Hz, 2H), 3.07 (q, J = 7.2 Hz, 2H), 1.43 (t, J = 7.2 Hz, 3H), 1.26 (t,

J = 7.2 Hz, 3H). Mass spectrum m/z 251.07 [M + H]⁺.

Ethyl 3-Ethyl-5-nitro-1H-indole-2-carboxylate (15b). The

compound was synthesized as per the general procedure as a pale

1

white solid (57% yield). R_f= 0.35 (EtOAc/hexanes = 20/80). H

Ethyl-2-acetyl Pentanoate (12a). To a ﬂask containing 300 mL

of anhydrous ethanol at 10 °C was added sodium metal (6.0 g, 261.0

mmol) portionwise under an argon atmosphere, and the mixture was

stirred for 30 min to complete dissolution. To this was added ethyl

acetoacetate (34.0 g, 261.0 mmol), and the resulting solution was

reﬂuxed for 30 min and allowed to cool to room temperature. This was

followed by addition of iodopropane (11a, 44.44 g, 261.0 mmol) over

NMR (400 MHz, chloroform-d): δ 9.02 (s, 1H), 8.69 (d, J = 2.0 Hz,

1H), 8.22 (dd, J = 8.8 Hz, J = 2.4 Hz, 1H), 7.43 (d, J = 9.6 Hz, 1H),

4.46 (q, J = 7.6 Hz, 2H), 3.16 (q, J = 7.2 Hz, 2H), 1.45 (t, J = 7.6 Hz,

3H), 1.31 (t, J = 7.2 Hz, 3H). Mass spectrum m/z 263.11 [M + H]⁺.

Ethyl 5-Chloro-3-(2-(1,3-dioxoisoindolin-2-yl)ethyl)-1H-in-

dole-2-carboxylate (15c). The compound was synthesized as per

1

the general procedure as a white solid (71% yield). H NMR (500

L

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

MHz, DMSO-d₆): δ 11.73 (s, 1H), 7.79 (s, 4H), 7.63 (d, J = 1.5 Hz,

1H), 7.38 (d, J = 8.5 Hz, 1H), 7.20 (dd, J = 8.0 Hz, 1.5 Hz, 1H), 4.19

(q, J = 7.0 Hz, 2H), 3.83 (t, J = 6.0 Hz, 2H), 3.35 (t, J = 6.0 Hz, 2H),

1.27 (t, J = 6.5 Hz, 3H). Mass spectrum m/z 397.09 [M + H]⁺

5-Chloro-3-ethyl-1H-indole-2-carboxylic Acid (16a). To a

solution of 15a (330 mg, 1.31 mmol) in 30 mL of dioxane was

added a solution of KOH (440 mg, 7.7 mol) in 3 mL of water, and the

resulting solution was reﬂuxed for 2 h. It was then cooled to room

temperature, concentrated under reduced pressure, and neutralized by

addition of 1 N hydrochloric acid. The precipitated acid was ﬁltered,

washed with cold water, and air-dried to give pure acid 16a (306 mg,

98% yield) as white solid. ¹H NMR (500 MHz, DMSO-d₆): δ 13.04 (s,

1H), 11.57 (s, 1H,), 7.71 (s, 1H), 7.40 (d, J = 9.0 Hz, 1H), 7.23 (dd, J

= 9.0 Hz, 2.0 Hz, 1H), 3.03 (q, J = 7.5 Hz, 2H), 1.16 (t, J = 7.5 Hz,

3H). Mass spectrum m/z 224.04 [M + H]⁺.

3-Ethyl-5-nitro-1H-indole-2-carboxylic Acid (16b). The com-

pound was synthesized as per the procedure for 16a, as a solid (72%

yield). ¹H NMR (400 MHz, DMSO-d₆): δ 12.14 (s, 1H), 8.67 (d, J =

2.0 Hz, 1H), 8.11 (dd, J = 8.8 Hz, 2.5 Hz, 1H), 7.54 (d, J = 8.8 Hz,

1H), 3.12 (q, J = 8.0 Hz, 2H), 1.22 (t, J = 7.6 Hz, 3H). Mass spectrum

m/z 235.06 [M + H]⁺.

3-Ethyl-5-nitro-N-[4-(piperidin-1-yl) phenethyl]-1H-indole-2-

carboxamide (17). The compound was synthesized as per the

procedure for 1, as a solid (82% yield). R_f= 0.7 (EtOAc/hexanes =

50/50). ¹H NMR (500 MHz, chloroform-d): δ 9.97 (s, 1H), 8.58 (d, J

= 2.0 Hz, 1H), 8.16 (dd, J = 9.5 Hz, J = 2.0 Hz, 1H), 7.44 (d, J = 9.5

Hz, 1H), 7.15 (d, J = 8.5 Hz, 2H), 6.93 (d, J = 8.5 Hz, 2H), 6.18−6.07

(m, 1H), 3.82 (q, J = 6.0 Hz, 2H), 3.14 (t, J = 5.5 Hz, 4H), 2.91 (t, J =

7.0 Hz, 2H), 2.79 (q, J = 7.5 Hz, 2H), 1.76−1.68 (m, 4H), 1.62−1.54

(m, 2H), 1.13 (t, J = 7.5 Hz, 3H). Mass spectrum m/z 421.22 [M +

H]⁺.

temperature for 15 min. It was quenched with cold water and extracted

with dichloromethane (3×), and the combined organic layer was

washed with brine, dried on Na₂SO₄, and evaporated under vacuum.

The resultant residue was puriﬁed on silica gel (5%−25%;

EtOAc:hexanes) to give pure compound 20 (388 mg, 87% yield). R_f

1

= 0.35 (EtOAc/hexanes = 20/80). H NMR (400 MHz, chloroform-

d): δ 9.79 (s, 1H), 7.46 (s, 1H), 7.35 (d, J = 8.8 Hz, 1H), 7.17−7.09

(m, 3H, especially 7.14, d, J = 8.0 Hz, 2H), 6.92 (d, J = 8.0 Hz, 2H),

6.03 (br t, J = 6.4 Hz, 1H), 3.80 (q, J = 6.0 Hz, 2H), 3.13 (br t, J = 5.6

Hz, 4H), 2.90 (t, J = 6.4 Hz, 2H), 2.70 (q, J = 7.6 Hz, 2H), 1.76−1.67

(m, 4H), 1.62−1.54 (m, 2H), 1.08 (t, J = 7.6 Hz, 3H). Mass spectrum

m/z 433.21 [M + H]⁺.

3-(2-Aminoethyl)-5-chloro-1H-indole-2-carboxylic Acid (21).

To a solution of 15c (10.0 g, 25.2 mmol) in 200 mL of ethanol was

added ethanolamine (3.08 g, 50.4 mmol), and the reaction mixture was

reﬂuxed for 14 h. It was then cooled to room temperature, volatiles

were removed under vacuum, and the mixture was partitioned in ethyl

acetate and water. The organic layer was separated, the aqueous layer

was extracted with ethyl acetate (3×), and the combined organic layer

was washed with brine and dried over Na₂SO₄. The solvent was

removed under vacuum to give the lactam intermediate as a pure white

1

solid (4.9 mg, 88% yield). R_f= 0.2 (EtOAc/hexanes = 50/50). H

NMR (500 MHz, DMSO-d ) δ 11.81 (s, 1H), 7.68 (d, J = 2.0 Hz,

6

1H), 7.66 (s, 1H), 7.39 (d, J = 9.0 Hz, 1H), 7.21 (dd, J = 7.0 Hz, 1.5

Hz, 1H), 3.54 (t, J = 4.0 Hz, 2H), 2.92 (t, J = 4.0 Hz, 2H). Mass

spectrum m/z 221.04 [M + H]⁺.

To this intermediate (4.5 g, 20.39 mmol) in 100 mL of

dioxane:H₂O (4:1) was added KOH (6.87 g, 122.0 mmol) in excess,

and the mixture was reﬂuxed overnight. It was then cooled to room

temperature, and the volatiles were removed under vacuum. The

residue was diluted with ice cold water and acidiﬁed to pH 5 with

concd hydrochloric acid to give a precipitate which was ﬁltered and air-

dried to give the desired 21 as a white solid. (4.77 g, 98% yield). R_f=

5-Amino-3-ethyl-N-[4-(piperidin-1-yl)phenethyl]-1H-indole-

2-carboxamide (18). To a solution of 17 (600 mg, 1.43 mmol) in 30

mL of anhydrous THF and anhydrous methanol (13:1) was added

NiCl₂·6H₂O (356 mg, 1.49 mmol) under an argon atmosphere, and

reaction mixture was stirred at room temperature for 45 min. The

mixture was then cooled to −5 °C to which was added NaBH₄(324

mg, 8.56 mmol) portionwise, and the reaction was gradually warmed

to room temperature while stirring for 1 h. Reaction was quenched

with saturated NH₄Cl and concentrated under reduced pressure. The

residue was diluted with ethyl acetate and water and ﬁltered, the

organic layer was separated, and the aqueous layer was extracted with

ethyl acetate (3×). The combined organic layer was washed with water

and brine, dried (Na₂SO₄), and evaporated under vacuum to yield 18

1

0.15 (MeOH/DCM = 20/80). H NMR (500 MHz, DMSO-d₆) δ

11.19 (s, 1H), 8.79 (br s, 3H), 7.62 (d, J = 2.0 Hz, 1H), 7.33 (d, J = 8.5

Hz, 1H), 7.08 (dd, J = 8.5 Hz, J = 2.0 Hz, 1H), 3.18 (t, J = 6.0 Hz,

2H), 3.01 (t, J = 6.0 Hz, 2H). Mass spectrum m/z 239.05 (M + H)⁺.

3-(2-((tert-Butoxycarbonyl)amino)ethyl)-5-chloro-1H-indole-

2-carboxylic Acid (22). To a solution of 21 (4.0 g, 16.76 mmol) in

80 mL of THF was added Boc anhydride (3.84 g, 17.60 mmol) at 0

°C. To this were added 50 mL of aq saturated NaHCO₃solution and

water (2:1), and the reaction mixture was stirred at 0 °C for 3 h and

then allowed to warm up to room temperature and stirred for 24 h.

Solvent was then removed under vacuum, ice cold water was added to

the residue, and it was acidiﬁed to pH 5 with cold 5% aq hydrochloric

acid. The resultant precipitate was ﬁltered, and the residue was washed

with cold water and air-dried to give crude product as a cream colored

solid which was recrystallized in methanol to give pure desired product

1

(479 mg, 86% yield). R_f= 0.8 (MeOH/DCM = 20/80). H NMR

(500 MHz, DMSO-d₆): δ 10.60 (s, 1H), 7.74 (t, J = 5.5 Hz, 1H), 7.08

(d, J = 9.0 Hz, 3H), 6.85 (d, J = 9.0 Hz, 2H), 6.69 (d, J = 2.0 Hz, 1H),

6.62 (dd, J = 9.0 Hz, 2.0 Hz, 1H), 4.56 (br s, 2H), 3.44 (q, J = 6.5 Hz,

2H), 3.06 (t, J = 5.5 Hz, 4H), 2.90 (q, J = 7.5 Hz, 2H), 2.74 (t, J = 7.5

Hz, 2H), 1.64−1.56 (m, 4H), 1.54−1.46 (m, 2H), 1.11 (t, J = 7.5 Hz,

3H). Mass spectrum m/z 390.24 [M + H]⁺.

5-Azido-3-ethyl-N-[4-(piperidin-1-yl)phenethyl]-1H-indole-

2-carboxamide (19). To a solution of 18 (400 mg, 1.10 mmol) in 40

mL of THF under argon atmosphere was added tert-butyl nitrite (1.6

g, 15.5 mmol) and TMSN₃(1.2 g, 10.42 mmol), and the reaction was

stirred overnight at room temperature. Solvent was evaporated under

reduced pressure to give crude product which was puriﬁed using ﬂash

column chromatography on silica gel (10%−40%; EtOAc:hexanes) to

yield pure 19 as a brown solid (180 mg, 42.0% yield). R_f= 0.81

(MeOH/DCM = 20/80). ¹H NMR (500 MHz, chloroform-d): δ 9.14

(s, 1H), 7.35 (d, J = 9.0 Hz, 1H), 7.20 (d, J = 2.0 Hz, 1H), 7.14 (d, J =

9.0 Hz, 2H), 6.95 (dd, J = 9.0 Hz, J = 2.0 Hz, 1H), 6.92 (d, J = 9.0 Hz,

2H), 5.98 (br t, J = 6.0 Hz, 1H,), 3.78 (q, J = 6.5 Hz, 2H), 3.13 (t, J =

5.5 Hz, 4H), 2.89 (t, J = 6.5 Hz, 2H), 2.70 (q, J = 8.0 Hz, 2H), 1.75−

1.68 (m, 4H), 1.64−1.54 (m, 2H), 1.08 (t, J = 8.0 Hz, 3H). Mass

spectrum m/z 417.23 [M + H]⁺.

1

22 (5.0 g, 88% yield). R_f= 0.25 (MeOH/DCM = 10/90). H NMR

(500 MHz, DMSO-d₆) δ 11.59 (s, 1H), 7.68 (s, 1H), 7.39 (d, J = 8.5

Hz, 1H), 7.20 (dd, J = 8.5 Hz, J = 1.5 Hz, 1H), 6.87 (t as br s, 1H),

3.16 (t, J = 8.5 Hz, 2H), 3.15 (t, J = 8.5 Hz, 2H), 1.31 (s, 9H). Mass

spectrum m/z 339.11 [M + H]⁺.

tert-Butyl (2-(5-Chloro-2-((4-(piperidin-1-yl)phenethyl)-

carbamoyl)-1H-indol-3-yl)ethyl)carbamate (23). The compound

was synthesized by coupling 22 with 8 as per the procedure for 1, as a

white solid (68% yield). R_f= 0.8 (EtOAc/hexanes = 50/50). ¹H NMR

(500 MHz, chloroform-d) δ 9.39 (s, 1H), 7.64 (s, 1H), 7.54 (d, J = 2.0

Hz, 1H), 7.32 (d, J = 8.5 Hz, 1H), 7.21 (dd, J = 8.5 Hz, 2.0 Hz, 1H),

7.15 (d, J = 8.0 Hz, 2H), 6.88 (d, J = 8.5 Hz, 2H), 4.92 (br s, 1H), 3.75

(dd, J = 15.5 Hz, 6.5 Hz, 2H), 3.21 (dd, J = 16.0 Hz, 6.0 Hz, 2H), 3.11

(t, J = 6.0 Hz, 4H), 3.09−3.03 (m, 2H), 2.95 (t, J = 8.0 Hz, 2H), 1.70

(quint, J = 5.5 Hz, 4H), 1.58−1.54 (m, 2H), 1.49 (s, 9H). Mass

spectrum m/z 526.25 [M + H]⁺.

3-(2-Aminoethyl)-5-chloro-N-(4-(piperidin-1-yl)phenethyl)-

1H-indole-2-carboxamide (24). To 100 mL of CH₂Cl₂was added

23 (2.0 g, 3.81 mmol) followed by dropwise addition of 10 mL of

TFA, and the reaction was stirred at room temperature for 3 h.

Volatiles were then removed under vacuum, and the crude product

was washed with saturated NaHCO₃and extracted in dichloromethane

3-Ethyl-5-isothiocyanato-N-[4-(piperidin-1-yl)phenethyl]-

1H-indole-2-carboxamide (20). To a solution of 18 (400 mg, 1.10

mmol) in 5 mL of CH₂Cl₂was added di(2-pyridyl) thionocarbonate

(308 mg, 1.32 mmol), and the reaction mixture was stirred at room

M

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

(3×). The combined organic layer was washed with brine, dried over

Na₂SO₄, and evaporated under vacuum to give 24 as a white solid

(1.47 g, 91% yield). R_f= 0.2 (MeOH/DCM = 20/80). ¹H NMR (500

MHz, chloroform-d) δ 10.32 (t as br s, 1H), 9.85 (s, 1H), 7.47 (d, J =

2.0 Hz, 1H), 7.34 (d, J = 8.5 Hz, 1H), 7.18 (dd, J = 9.0 Hz, 2.0 Hz,

1H), 7.13 (d, J = 9.0 Hz, 2H), 6.89 (d, J = 8.5 Hz, 2H), 3.72 (q, J = 6.5

Hz, 2H), 3.12 (t, J = 5.5 Hz, 4H), 2.94−2.84 (m, 6H), 1.76−1.66 (m,

4H), 1.62−1.52 (m, 2H), 1.35 (s, 2H). Mass spectrum m/z 426.20 [M

+ H]⁺.

3-(2-Azidoethyl)-5-chloro-N-(4-(piperidin-1-yl)phenethyl)-

1H-indole-2-carboxamide (25). To a 10 mL solution of NaN₃(2.07

g, 31.9 mmol) in H₂O was added triﬂuoromethanesulfonyl anhydride

(3.0 g, 10.63 mmol) in DCM at 0 °C and stirred for 2 h while

maintaining the temperature. The organic layer was separated, the

aqueous layer was extracted with DCM (2×), and the organic layers

were combined to aﬀord TfN₃. In a separate round-bottom ﬂask, a

solution of 24 (1.0 g, 2.35 mmol) in 20 mL of H₂O:methanol (1:20)

was treated with K₂CO₃(2.6 g, 18.83 mmol) and CuSO₄(751 mg,

4.71 mmol). To this mixture was added the above TfN₃solution, and

it was stirred at room temperature for 18 h. Volatiles were then

removed under vacuum, and the residue was dissolved in DCM,

washed with water and brine, and dried over Na₂SO₄. The organic

layer was concentrated under vacuum, and the crude product was

puriﬁed on silica gel (0%−20%; EtOAc:hexanes) to obtain 25 as a

pure compound (679 mg, 64%). R_f= 0.7 (EtOAc/hexanes = 50/50).

¹H NMR (500 MHz, chloroform-d) δ 11.49 (s, 1H), 8.16−8.06 (m,

yield) as a yellow solid. ¹H NMR (500 MHz, DMSO-d₆) δ: 8.57 (t, J =

1.5 Hz, 1H), 8.50 (td, J = 8.0 Hz, J = 1.5 Hz, 1H), 8.23 (ddd, J = 8.0

Hz, J = 2.5 Hz, J = 1.0 Hz, 1H), 7.74 (t, J = 8.0 Hz, 1H), 7.63 (dd, J =

8.0 Hz, J = 7.0 Hz, 1H), 7.28 (d, J = 7.0 Hz, 1H), 6.51 (d, J = 8.0 Hz,

1H), 3.49 (t, J = 6.5 Hz, 4H), 2.04−1.94 (quin, J = 3.0 Hz, 4H). Mass

spectrum m/z 270.1 [M + H]⁺.

3-(6-(Pyrrolidin-1-yl)pyridin-2-yl)aniline (31). To a solution of

30 (5 g, 18.57 mmol) in methanol (100 mL) was added catalytic

amounts of Raney-nickel, and the resultant mixture was stirred for 3 h

under hydrogen atmosphere. Raney-nickel was ﬁltered oﬀ and, ﬁltrate

was concentrated and puriﬁed by silica gel chromatography (20−70%;

1

EtOAc:hexanes) to give 31 (4.25 g, 96% yield) as a whitish solid. H

NMR (500 MHz, chloroform-d) δ: 7.52−7.48 (m, 1H), 7.46 (t, J = 2.0

Hz, 1H), 7.45−7.41 (m, 1H), 7.29 (s, 1H), 7.23 (t, J = 8.0 Hz, 1H),

6.99 (d, J = 7.0 Hz, 1H), 6.72 (ddd, J = 5.5 Hz, J = 2.5 Hz, J = 1.0 Hz,

1H), 3.75 (br s, 2H), 3.57 (t, J = 6.5 Hz, 4H), 2.08−1.80 (m, 4H).

Mass spectrum m/z 240.2 [M + H]⁺.

1-(4-Azidophenyl)-3-(3-(6-(pyrrolidin-1-yl)pyridin-2-yl)-

phenyl)urea (33). To a solution of triphosgene (70 mg, 0.21 mmol)

in toluene (10 mL) was added 31 (130 mg, 0.54 mmol) followed by

Et₃N (2.4 mL, 23.6 mmol) under inert atmosphere, and the mixture

was heated to 70 °C for 3 h under argon atmosphere. The reaction was

concentrated under reduced pressure and carried forward to the next

step without any puriﬁcation. To a solution of this isocyanate

intermediate in DCM (4 mL) was added 4-azidoaniline 32 (72 mg,

0.167 mmol) followed by Et₃N (2.4 mL, 23.6 mmol), and the mixture

was stirred at 0 °C for 6 h. The reaction mixture was diluted in 20 mL

of DCM and water (1:1). The organic layer was separated, washed

with water, dried (Na₂SO₄), and concentrated under reduced pressure.

The crude product was puriﬁed by silica gel chromatography (10−

50%; EtOAc:hexanes) to aﬀord the desired product 33 (150 mg, 75%

1H), 7.75 (s, 1H), 7.43 (d, J = 8.5 Hz, 1H), 7.21 (d, J = 8.5 Hz, 1H),

7.09 (d, J = 8.5 Hz, 2H), 6.86 (d, J = 8.0 Hz, 2H), 3.56−3.42 (m, 4H),

3.24 (t, J = 7.0 Hz, 2H), 3.13−3.01 (m, 4H), 2.76 (t, J = 7.0 Hz, 2H),

1.67−1.55 (m, 4H), 1.55−1.44 (m, 2H). Mass spectrum m/z 452.19

[M + H]⁺.

1

5-Chloro-3-(2-isothiocyanatoethyl)-N-(4-(piperidin-1-yl)-

phenethyl)-1H-indole-2-carboxamide (26). To a solution of 24

(1.0 g, 2.353 mmol) in 50 mL of CH₂Cl₂was added di(2-pyridyl)

thionocarbonate (656 mg, 2.82 mmol), and the reaction mixture was

stirred at room temperature for 15 min. It was quenched with cold

water and extracted with dichloromethane (3×), and the combined

organic layer was washed with brine, dried on Na₂SO₄, and evaporated

under vacuum. The resultant residue was puriﬁed on silica gel (5%−

25%; EtOAc:hexanes) to give pure compound 26 (956 mg, 87%

yield) as white solid. H NMR (500 MHz, DMSO-d₆) δ 8.78 (s, 1H),

8.75 (s, 1H), 8.07 (t, J = 2.0 Hz, 1H), 7.63 (dt, J = 7.5 Hz, 1.5 Hz,

1H), 7.56 (dd, J = 8.5 Hz, 7.0 Hz, 2H), 7.52 (d, J = 9.0 Hz, 2H) 7.34

(t, J = 7.5 Hz, 1H), 7.08−7.03 (m, 3H), 6.42 (d, J = 9.0 Hz, 1H),

3.54−3.42 (m, 4H), 2.04−1.92 (m, 4H). Mass spectrum m/z 400.18

[M + H]⁺.

1-(4-Isothiocyanatophenyl)-3-(3-(6-(pyrrolidin-1-yl)pyridin-

2-yl)phenyl)urea (34). To a solution of 33 (25 mg, 0.06 mmol) in 5

mL of benzene under an argon atmosphere was added triphenylphos-

phine (34.2 mg, 0.12 mmol), and the reaction mixture was reﬂuxed for

4 h. The reaction mixture was cooled to room temperature, 1 mL of

CS₂was added to this, and the reaction mix was stirred at 40 °C for 12

h. Volatiles were evaporated under reduced pressure to obtain crude

product which was puriﬁed using ﬂash column chromatography

(10%−40%; EtOAc:hexanes) to obtain pure 34 as a white solid (17

1

yield). R_f= 0.35 (EtOAc/hexanes = 20/80). H NMR (500 MHz,

DMSO-d₆) δ 11.52 (s, 1H), 8.17 (t, J = 5.5 Hz, 1H), 7.81 (d, J = 2.0

Hz, 1H), 7.44 (d, J = 8.5 Hz, 1H), 7.22 (ddd, J = 8.5 Hz, 2.0 Hz, 1.0

Hz, 1H), 7.10 (d, J = 8.5 Hz, 2H), 6.86 (d, J = 8.5 Hz, 2H), 3.82 (t, J =

7.0 Hz, 2H), 3.48 (q, J = 7.0 Hz, 2H), 3.38 (t, J = 7.0 Hz, 2H), 3.07 (t,

J = 5.5 Hz, 4H), 2.76 (t, J = 7.0 Hz, 2H), 1.65−1.56 (m, 4H), 1.56−

1.46 (m, 2H). Mass spectrum m/z 468.16 [M + H]⁺.

1

mg, 74% yield). R_f= 0.78 (MeOH/DCM = 20/80). H NMR (500

2-Bromo-6-(pyrrolidin-1-yl)pyridine (29). A mixture of 27

(10.0 g, 42.2 mmol) and 28 (13.87 mL, 169 mmol) was stirred for

1 h at room temperature. The reaction mixture was quenched with 100

mL of saturated NaHCO₃solution and diluted with 100 mL of

dichloromethane. The organic layer was separated, washed with water

and brine, and dried (MgSO₄) and concentrated under reduced

pressure. The resulting crude product was crystallized from methanol

(50 mL) to aﬀord the desired product 29 (8.5 g, 37.4 mmol, 89%

yield) as a white solid. ¹H NMR (500 MHz, methanol-d₄) δ: 7.22 (t, J

= 8.0 Hz, 1H), 6.64 (d, J = 7.5 Hz, 1H), 6.23 (d, J = 8.0 Hz, 1H), 3.43

(t, J = 6.5 Hz, 4H), 2.04−1.94 (m, 4H). Mass spectrum m/z 228.01

[M + H]⁺.

2-(3-Nitrophenyl)-6-(pyrrolidin-1-yl)pyridine (30). To a sol-

ution of 29 (2 g, 8.81 mmol) and 3-nitrophenylboronic acid (1.62 g,

9.69 mmol) in 1,2-dimethoxyethane (10 mL) were added Ba(OH)₂

(3.32 g, 19.37 mmol) and water (4 mL) under argon atmosphere. The

contents were degassed, Pd(Ph₃P)₄(0.305 g, 0.26 mmol) was added,

and the resulting mixture was irradiated with microwaves at 150 °C for

15 min. It was then diluted with 200 mL of ethyl acetate and water

(1:1), and the organic layer was separated, washed with water and

brine, and dried over MgSO₄. Volatiles were concentrated under

reduced pressure to give crude product which was puriﬁed by silica gel

chromatography (20−70%; EtOAc:hexanes) to give 30 (2.05 g, 86%

MHz, DMSO-d₆) δ 8.96 (s, 1H), 8.84 (s, 1H), 8.07 (t, J = 2.0 Hz, 1H),

7.64 (d, J = 7.5 Hz, 1H), 7.59−7.51 (m, 4H, esp. 7.54, d, J = 9.5 Hz,

2H), 7.38 (d, J = 9.0 Hz, 2H), 7.35 (t, J = 7.0 Hz, 1H), 7.06 (d, J = 7.5

Hz, 1H), 6.42 (d, J = 8.5 Hz, 1H), 3.54−3.42 (m, 4H), 2.04−1.92 (m,

4H). Mass spectrum m/z 416.15 [M + H]⁺.

1-(4-Benzoylphenyl)-3-(3-(6-(pyrrolidin-1-yl)pyridin-2-yl)-

phenyl)urea (36). To a solution of triphosgene (23 mg, 0.77 mmol)

in toluene (2 mL) was added 35 (40 mg, 0.16 mmol) followed by

Et₃N (0.78 mL, 7.8 mmol) under inert atmosphere, and the mixture

was heated to 70 °C for 3 h under argon atmosphere. The reaction was

concentrated under reduced pressure and carried forward to the next

step without any puriﬁcation.

To this intermediate dissolved in THF (10 mL) was added 31 (39

mg, 0.167 mmol) followed by Et₃N (0.78 mL, 7.8 mmol), and the

mixture was stirred at 0 °C for 1 h and then at room temperature for 5

h. The reaction mixture was concentrated under reduced pressure, and

crude product was recrystallized from methanol to give desired

1

product 36 (60 mg, 78% yield) as a white solid. H NMR (500 MHz,

DMSO-d₆) δ 9.19 (s, 1H), 8.92 (s, 1H), 8.10 (t, J = 2.0 Hz, 1H), 7.75

(d, J = 9.0 Hz, 2H), 7.71 (dd, J = 8.0 Hz, 1.5 Hz, 2H), 7.69−7.63 (m,

4H), 7.61−7.53 (m, 4H), 7.37 (t, J = 8.0 Hz, 1H), 7.07 (d, J = 7.5 Hz,

1H), 6.43 (d, J = 8.5 Hz, 1H), 3.54−3.45 (m, 4H), 2.03−1.93 (m,

4H). Mass spectrum m/z 463.21 [M + H]⁺.

N

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

(2) Pacher, P.; Batkai, S.; Kunos, G. The Endocannabinoid System as

an Emerging Target of Pharmacotherapy. Pharmacol. Rev. 2006, 58,

389−462.

(3) Pacher, P.; Kunos, G. Modulating the Endocannabinoid System

in Human Health and Disease-Successes and Failures. FEBS J. 2013,

280, 1918−1943.

ASSOCIATED CONTENT

■

S

* Supporting Information

The Supporting Information is available free of charge on the

ACS Publications website at DOI: 10.1021/acs.jmed-

chem.5b01303.

(4) Maccarrone, M.; Bab, I.; Biro, T.; Cabral, G. A.; Dey, S. K.; Di

Marzo, V.; Konje, J. C.; Kunos, G.; Mechoulam, R.; Pacher, P.;

Sharkey, K. A.; Zimmer, A. Endocannabinoid Signaling at the

Periphery: 50 Years after THC. Trends Pharmacol. Sci. 2015, 36,

277−296.

(5) Devane, W. A.; Dysarz, F. A., III; Johnson, M. R.; Melvin, L. S.;

Howlett, A. C. Determination and Characterization of a Cannabinoid

Receptor in Rat Brain. Mol. Pharmacol. 1988, 34, 605−613.

(6) Matsuda, L. A.; Lolait, S. J.; Brownstein, M. J.; Young, A. C.;

Bonner, T. I. Structure of a Cannabinoid Receptor and Functional

Expression of the Cloned cDNA. Nature 1990, 346, 561−564.

(7) Munro, S.; Thomas, K. L.; Abu-Shaar, M. Molecular Character-

ization of a Peripheral Receptor for Cannabinoids. Nature 1993, 365,

61−65.

SMILES data (CSV)

AUTHOR INFORMATION

■

Corresponding Author

*E-mail: g.thakur@neu.edu. Phone: 617 373 8163. Fax: 617

373 8886.

Notes

The authors declare no competing ﬁnancial interest.

ACKNOWLEDGMENTS

■

The work was supported by National Institutes of Health grants

DA027113 and EY024717 to G.A.T. and DA09158 to A.M. A

portion of this work was submitted in 2011 by A. Kulkarni in

partial fulﬁllment of M.S. degree requirements from North-

eastern University, Boston, MA.

(8) Galiegue, S.; Mary, S.; Marchand, J.; Dussossoy, D.; Carriere, D.;

Carayon, P.; Bouaboula, M.; Shire, D.; Le Fur, G.; Casellas, P.

Expression of Central and Peripheral Cannabinoid Receptors in

Human Immune Tissues and Leukocyte Subpopulations. Eur. J.

Biochem. 1995, 232, 54−61.

(9) Cabral, G. A.; Raborn, E. S.; Griffin, L.; Dennis, J.; Marciano-

Cabral, F. CB2 Receptors in the Brain: Role in Central Immune

Function. Br. J. Pharmacol. 2008, 153, 240−251.

(10) Rahn, E. J.; Deng, L.; Thakur, G. A.; Vemuri, K.; Zvonok, A. M.;

Lai, Y. Y.; Makriyannis, A.; Hohmann, A. G. Prophylactic Cannabinoid

Administration Blocks the Development of Paclitaxel-Induced Neuro-

pathic Nociception during Analgesic Treatment and Following

Cessation of Drug Delivery. Mol. Pain 2014, 10, 27.

(11) Deng, L.; Guindon, J.; Vemuri, V. K.; Thakur, G. A.; White, F.

A.; Makriyannis, A.; Hohmann, A. G. The Maintenance of Cisplatin-

and Paclitaxel-Induced Mechanical and Cold Allodynia Is Suppressed

by Cannabinoid CB(2) Receptor Activation and Independent of

CXCR4 Signaling in Models of Chemotherapy-Induced Peripheral

Neuropathy. Mol. Pain 2012, 8, 71.

(12) Wilkerson, J. L.; Gentry, K. R.; Dengler, E. C.; Wallace, J. A.;

Kerwin, A. A.; Armijo, L. M.; Kuhn, M. N.; Thakur, G. A.;

Makriyannis, A.; Milligan, E. D. Intrathecal Cannabilactone CB(2)R

Agonist, AM1710, Controls Pathological Pain and Restores Basal

Cytokine Levels. Pain 2012, 153, 1091−1106.

(13) Rahn, E. J.; Thakur, G. A.; Wood, J. A.; Zvonok, A. M.;

Makriyannis, A.; Hohmann, A. G. Pharmacological Characterization of

AM1710, a Putative Cannabinoid CB2 Agonist from the Cannabi-

lactone Class: Antinociception Without Central Nervous System Side-

Effects. Pharmacol., Biochem. Behav. 2011, 98, 493−502.

(14) Khanolkar, A. D.; Lu, D.; Ibrahim, M.; Duclos, R. I., Jr.; Thakur,

G. A.; Malan, T. P., Jr.; Porreca, F.; Veerappan, V.; Tian, X.; George,

C.; Parrish, D. A.; Papahatjis, D. P.; Makriyannis, A. Cannabilactones:

A Novel Class of CB2 Selective Agonists with Peripheral Analgesic

Activity. J. Med. Chem. 2007, 50, 6493−6500.

(15) Wilkerson, J. L.; Gentry, K. R.; Dengler, E. C.; Wallace, J. A.;

Kerwin, A. A.; Kuhn, M. N.; Zvonok, A. M.; Thakur, G. A.;

Makriyannis, A.; Milligan, E. D. Immunofluorescent Spectral Analysis

Reveals the Intrathecal Cannabinoid Agonist, AM1241, Produces

Spinal Anti-Inflammatory Cytokine Responses in Neuropathic Rats

Exhibiting Relief from Allodynia. Brain Behav. 2012, 2, 155−177.

(16) Han, S.; Thatte, J.; Buzard, D. J.; Jones, R. M. Therapeutic

Utility of Cannabinoid Receptor Type 2 (CB(2)) Selective Agonists. J.

Med. Chem. 2013, 56, 8224−8256.

ABBREVIATIONS USED

■

AEA, arachidonoylethanol amine; 2-AG, 2-arachidonoylglycer-

ol; BSA, bovine serum albumin; cAMP, cyclic adenosine

monophosphate; CB1R, cannabinoid 1 receptor; CB2R,

cannabinoid 2 receptor; CHO, Chinese hamster ovary cells;

CI, conﬁdence interval; CO₂, carbon dioxide; CNS, central

nervous system; DMSO, dimethyl sulfoxide; DCM, dichloro-

methane; DIPEA, N,N-diisopropylethylamine; DPT, di-2-

pyridyl thionocarbonate; EDTA, ethylenediaminetetraacetic

acid; EtOAc, ethyl acetate; Et₃N, triethylamine; EDCI, 1-

ethyl-3-(3-(dimethylamino)propyl)carbodiimide; GDP, guano-

sine diphosphate; GTP, guanosine triphosphate; GPCRs, G-

protein-coupled receptors; [³⁵S]GTPγS, guanosine 5′-O-(3-

[³⁵S]thio)triphosphate; HEK293, human embryonic kidney

293 cells; HPLC, high performance liquid chromatography;

HTS, high-throughput screening; Hz, hertz; HOBT, hydrox-

ybenzotriazole; K₂CO₃, potassium carbonate; KOH, potassium

hydroxide; LAPS, ligand-assisted protein structure; LCMS,

liquid chromatography mass spectrometry; LiAlH₄, lithium

aluminum hydride; MgSO₄, magnesium sulfate; MHz, mega-

hertz; MeOH, methanol; NaBH₄, sodium borohydride; NAMs,

negative allosteric modulators; Na₂SO₄, sodium sulfate;

NaOAc, sodium acetate; NaHCO₃, sodium bicarbonate;

NaNO₂, sodium nitrite; NiCl₂·6H₂O, nickel chloride hexahy-

drate; NH₄OAc, ammonium acetate; NH₄Cl, ammonium

chloride; NMR, nuclear magnetic resonance spectroscopy;

NMP, N-methylpyrrolidinone; OCC, optimized cell culture;

RLU, relative light units; THF, tetrahydrofuran; TLC, thin-

layer chromatography; SAR, structure−activity relationship;

SEM, standard error of mean; Tris, tris(hydroxymethyl)-

aminomethane; TMSN₃, trimethylsilyl azide; UV, ultraviolet

REFERENCES

■

(1) Pertwee, R. G.; Howlett, A. C.; Abood, M. E.; Alexander, S. P.; Di

Marzo, V.; Elphick, M. R.; Greasley, P. J.; Hansen, H. S.; Kunos, G.;

Mackie, K.; Mechoulam, R.; Ross, R. A. International Union of Basic

and Clinical Pharmacology. LXXIX. Cannabinoid Receptors and their

Ligands: Beyond CB(1) and CB(2). Pharmacol. Rev. 2010, 62, 588−

631.

(17) Dhopeshwarkar, A.; Mackie, K. CB2 Cannabinoid Receptors as

a Therapeutic Target-What Does the Future Hold? Mol. Pharmacol.

2014, 86, 430−437.

(18) Palmer, S. L.; Thakur, G. A.; Makriyannis, A. Cannabinergic

Ligands. Chem. Phys. Lipids 2002, 121, 3−19.

O

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

(19) Pertwee, R. G. Emerging Strategies for Exploiting Cannabinoid

Receptor Agonists as Medicines. Br. J. Pharmacol. 2009, 156, 397−411.

(20) Thakur, G. A.; Tichkule, R.; Bajaj, S.; Makriyannis, A. Latest

Advances in Cannabinoid Receptor Agonists. Expert Opin. Ther. Pat.

2009, 19, 1647−1673.

(21) Moreira, F. A.; Grieb, M.; Lutz, B. Central Side-Effects of

Therapies Based on CB1 Cannabinoid Receptor Agonists and

Antagonists: Focus on Anxiety and Depression. Best Pract. Res. Clin.

Endocrinol. Metab. 2009, 23, 133−144.

(22) Moreira, F. A.; Crippa, J. A. The Psychiatric Side-Effects of

Rimonabant. Rev. Bras. Psiquiatr. 2009, 31, 145−153.

(37) Ignatowska-Jankowska, B. M.; Baillie, G. L.; Kinsey, S.; Crowe,

M.; Ghosh, S.; Owens, R. A.; Damaj, I. M.; Poklis, J.; Wiley, J. L.;

Zanda, M.; Zanato, C.; Greig, I. R.; Lichtman, A. H.; Ross, R. A. A

Cannabinoid CB1 Receptor-Positive Allosteric Modulator Reduces

Neuropathic Pain in the Mouse with No Psychoactive Effects.

Neuropsychopharmacology 2015, DOI: 10.1038/npp.2015.148.

(38) Gamage, T. F.; Ignatowska-Jankowska, B. M.; Wiley, J. L.;

Abdelrahman, M.; Trembleau, L.; Greig, I. R.; Thakur, G. A.; Tichkule,

R.; Poklis, J.; Ross, R. A.; Pertwee, R. G.; Lichtman, A. H. In-vivo

Pharmacological Evaluation of the CB1-Receptor Allosteric Modulator

Org-27569. Behav. Pharmacol. 2014, 25, 182−185.

(23) Janero, D. R. Cannabinoid-1 Receptor (CB1R) Blockers as

Medicines: Beyond Obesity and Cardiometabolic Disorders to

Substance Abuse/Drug Addiction with CB1R Neutral Antagonists.

Expert Opin. Emerging Drugs 2012, 17, 17−29.

(24) Kunos, G.; Osei-Hyiaman, D.; Batkai, S.; Sharkey, K. A.;

Makriyannis, A. Should Peripheral CB(1) Cannabinoid Receptors Be

Selectively Targeted for Therapeutic Gain? Trends Pharmacol. Sci.

2009, 30, 1−7.

(25) Bergman, J.; Delatte, M. S.; Paronis, C. A.; Vemuri, K.; Thakur,

G. A.; Makriyannis, A. Some Effects of CB1 Antagonists with Inverse

Agonist and Neutral Biochemical Properties. Physiol. Behav. 2008, 93,

666−670.

(39) Ding, Y.; Qiu, Y.; Jing, L.; Thorn, D. A.; Zhang, Y.; Li, J. X.

Behavioral Effects of the Cannabinoid CB1 Receptor Allosteric

Modulator ORG27569 in Rats. Pharmacol. Res. Perspect. 2014, 2 (6),

e00069.

(40) Thakur, G. A.; Tichkule, R. B.; Kulkarni, P. M.; Kulkarni, A. R.

Allosteric modulators of the cannabinoid 1 receptor. WO/2015/

027160A2, Aug 22, 2014.

(41) Baillie, G. L.; Horswill, J. G.; Anavi-Goffer, S.; Reggio, P. H.;

Bolognini, D.; Abood, M. E.; McAllister, S.; Strange, P. G.; Stephens,

G. J.; Pertwee, R. G.; Ross, R. A. CB(1) Receptor Allosteric

Modulators Display Both Agonist and Signaling Pathway Specificity.

Mol. Pharmacol. 2013, 83, 322−338.

(26) Price, M. R.; Baillie, G. L.; Thomas, A.; Stevenson, L. A.; Easson,

M.; Goodwin, R.; McLean, A.; McIntosh, L.; Goodwin, G.; Walker, G.;

Westwood, P.; Marrs, J.; Thomson, F.; Cowley, P.; Christopoulos, A.;

Pertwee, R. G.; Ross, R. A. Allosteric Modulation of the Cannabinoid

CB1 receptor. Mol. Pharmacol. 2005, 68, 1484−1495.

(27) Horswill, J. G.; Bali, U.; Shaaban, S.; Keily, J. F.; Jeevaratnam, P.;

Babbs, A. J.; Reynet, C.; Wong-Kai-In, P. PSNCBAM-1, A Novel

Allosteric Antagonist at Cannabinoid CB1 Receptors with Hypophagic

Effects in Rats. Br. J. Pharmacol. 2007, 152, 805−814.

(42) Shore, D. M.; Baillie, G. L.; Hurst, D. H.; Navas, F., 3rd;

Seltzman, H. H.; Marcu, J. P.; Abood, M. E.; Ross, R. A.; Reggio, P. H.

Allosteric Modulation of a Cannabinoid G Protein-Coupled Receptor:

Binding Site Elucidation and Relationship to G Protein Signaling. J.

Biol. Chem. 2014, 289, 5828−5845.

(43) Kapur, A.; Samaniego, P.; Thakur, G. A.; Makriyannis, A.;

Abood, M. E. Mapping the Structural Requirements in the CB1

Cannabinoid Receptor Transmembrane Helix II for Signal Trans-

duction. J. Pharmacol. Exp. Ther. 2008, 325, 341−348.

(28) Pamplona, F. A.; Ferreira, J.; Menezes de Lima, O., Jr.; Duarte,

F. S.; Bento, A. F.; Forner, S.; Villarinho, J. G.; Bellocchio, L.; Wotjak,

C. T.; Lerner, R.; Monory, K.; Lutz, B.; Canetti, C.; Matias, I.; Calixto,

J. B.; Marsicano, G.; Guimaraes, M. Z.; Takahashi, R. N. Anti-

Inflammatory Lipoxin A4 is an Endogenous Allosteric Enhancer of

CB1 Cannabinoid Receptor. Proc. Natl. Acad. Sci. U. S. A. 2012, 109,

21134−21139.

(29) Vallee, M.; Vitiello, S.; Bellocchio, L.; Hebert-Chatelain, E.;

Monlezun, S.; Martin-Garcia, E.; Kasanetz, F.; Baillie, G. L.; Panin, F.;

Cathala, A.; Roullot-Lacarriere, V.; Fabre, S.; Hurst, D. P.; Lynch, D.

L.; Shore, D. M.; Deroche-Gamonet, V.; Spampinato, U.; Revest, J. M.;

Maldonado, R.; Reggio, P. H.; Ross, R. A.; Marsicano, G.; Piazza, P. V.

Pregnenolone Can Protect the Brain from Cannabis Intoxication.

Science 2014, 343, 94−98.

(44) Ahn, K. H.; Mahmoud, M. M.; Kendall, D. A. Allosteric

Modulator ORG27569 Induces CB1 Cannabinoid Receptor High

Affinity Agonist Binding State, Receptor Internalization, and G_i

Protein-Independent ERK1/2 Kinase Activation. J. Biol. Chem. 2012,

287, 12070−12082.

(45) Pei, Y.; Mercier, R. W.; Anday, J. K.; Thakur, G. A.; Zvonok, A.

M.; Hurst, D.; Reggio, P. H.; Janero, D. R.; Makriyannis, A. Ligand-

Binding Architecture of Human CB2 Cannabinoid Receptor: Evidence

for Receptor Subtype-Specific Binding Motif and Modeling GPCR

Activation. Chem. Biol. 2008, 15, 1207−1219.

(46) Picone, R. P.; Khanolkar, A. D.; Xu, W.; Ayotte, L. A.; Thakur,

G. A.; Hurst, D. P.; Abood, M. E.; Reggio, P. H.; Fournier, D. J.;

Makriyannis, A. (−)-7′-Isothiocyanato-11-hydroxy-1′,1′-dimethylhep-

tylhexahydrocannabinol (AM841), a High-Affinity Electrophilic

Ligand, Interacts Covalently with a Cysteine in Helix Six and Activates

the CB1 Cannabinoid Receptor. Mol. Pharmacol. 2005, 68, 1623−

1635.

(30) Wang, C. I.; Lewis, R. J. Emerging Opportunities for Allosteric

Modulation of G-Protein Coupled Receptors. Biochem. Pharmacol.

2013, 85, 153−162.

(31) Fay, J. F.; Farrens, D. L. A Key Agonist-Induced Conformational

Change in the Cannabinoid Receptor CB1 Is Blocked by the Allosteric

Ligand Org 27569. J. Biol. Chem. 2012, 287, 33873−33882.

(32) Conn, P. J.; Christopoulos, A.; Lindsley, C. W. Allosteric

Modulators of GPCRs: A Novel Approach for the Treatment of CNS

Disorders. Nat. Rev. Drug Discovery 2009, 8, 41−54.

(33) Navarro, H. A.; Howard, J. L.; Pollard, G. T.; Carroll, F. I.

Positive Allosteric Modulation of the Human Cannabinoid (CB)

Receptor by RTI-371, A Selective Inhibitor of the Dopamine

Transporter. Br. J. Pharmacol. 2009, 156, 1178−1184.

(34) Priestley, R. S.; Nickolls, S. A.; Alexander, S. P.; Kendall, D. A. A

Potential Role for Cannabinoid Receptors in the Therapeutic Action

of Fenofibrate. FASEB J. 2015, 29, 1446−1455.

(35) Laprairie, R. B.; Bagher, A. M.; Kelly, M. E.; Denovan-Wright, E.

M. Cannabidiol is a Negative Allosteric Modulator of the Type 1

Cannabinoid Receptor. Br. J. Pharmacol. 2015, 172, 4790.

(36) Thakur, G. A.; Kulkarni, P. M. Preparation of Phenylindole

Derivatives for Use as CB1 Cannabinoid Receptor Modulators.

WO2013103967A1, Jul 11, 2013.

(47) Janero, D. R.; Yaddanapudi, S.; Zvonok, N.; Subramanian, K. V.;

Shukla, V. G.; Stahl, E.; Zhou, L.; Hurst, D.; Wager-Miller, J.; Bohn, L.

M.; Reggio, P. H.; Mackie, K.; Makriyannis, A. Molecular-Interaction

and Signaling Profiles of AM3677, a Novel Covalent Agonist Selective

for the Cannabinoid 1 Receptor. ACS Chem. Neurosci. 2015, 6, 1400.

(48) Szymanski, D. W.; Papanastasiou, M.; Melchior, K.; Zvonok, N.;

Mercier, R. W.; Janero, D. R.; Thakur, G. A.; Cha, S.; Wu, B.; Karger,

B.; Makriyannis, A. Mass Spectrometry-Based Proteomics of Human

Cannabinoid Receptor 2: Covalent Cysteine 6.47(257)-Ligand

Interaction Affording Megagonist Receptor Activation. J. Proteome.

Res. 2011, 10, 4789−4798.

(49) Makriyannis, A. 2012 Division of Medicinal Chemistry Award

Address. Trekking the Cannabinoid Road: A Personal Perspective. J.

Med. Chem. 2014, 57, 3891−3911.

(50) Weichert, D.; Gmeiner, P. Covalent Molecular Probes for Class

A G Protein-Coupled Receptors: Advances and Applications. ACS

Chem. Biol. 2015, 10, 1376−1386.

(51) Davie, B. J.; Sexton, P. M.; Capuano, B.; Christopoulos, A.;

Scammells, P. J. Development of a Photoactivatable Allosteric Ligand

P

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

Article

for the M1 Muscarinic Acetylcholine Receptor. ACS Chem. Neurosci.

2014, 5, 902−907.

(68) Isoda, T.; Hayashi, K.; Tamai, S.; Kumagai, T.; Nagao, Y.

Efficient Synthesis of Isothiocyanates Based on the Tandem

Staudinger/Aza-Wittig Reactions and Mechanistic Consideration of

the Tandem Reactions. Chem. Pharm. Bull. 2006, 54, 1616−1619.

(69) Chen, L.; Jin, L.; Zhou, N. An Update of Novel Screening

Methods for GPCR in Drug Discovery. Expert Opin. Drug Discovery

2012, 7, 791−806.

(52) Davie, B. J.; Valant, C.; White, J. M.; Sexton, P. M.; Capuano, B.;

Christopoulos, A.; Scammells, P. J. Synthesis and Pharmacological

Evaluation of Analogues of Benzyl Quinolone Carboxylic Acid

(BQCA) Designed To Bind Irreversibly to an Allosteric Site of the

M(1) Muscarinic Acetylcholine Receptor. J. Med. Chem. 2014, 57,

5405−5418.

(70) Tian, X.; Kang, D. S.; Benovic, J. L. Beta-arrestins and G

Protein-Coupled Receptor Trafficking. Handb. Exp. Pharmacol. 2014,

219, 173−186.

(53) Pace, N. J.; Weerapana, E. Diverse Functional Roles of Reactive

Cysteines. ACS Chem. Biol. 2013, 8, 283−296.

(54) Tahtaoui, C.; Balestre, M. N.; Klotz, P.; Rognan, D.; Barberis,

C.; Mouillac, B.; Hibert, M. Identification of the Binding Sites of the

SR49059 Nonpeptide Antagonist into the V1a Vasopressin Receptor

Using Sulfydryl-Reactive Ligands and Cysteine Mutants as Chemical

Sensors. J. Biol. Chem. 2003, 278, 40010−40019.

(55) Keenan, C. M.; Storr, M. A.; Thakur, G. A.; Wood, J. T.; Wager-

Miller, J.; Straiker, A.; Eno, M. R.; Nikas, S. P.; Bashashati, M.; Hu, H.;

Mackie, K.; Makriyannis, A.; Sharkey, K. A. AM841, a Covalent

Cannabinoid Ligand, Powerfully Slows Gastrointestinal Motility in

Normal and Stressed Mice in a Peripherally Restricted Manner. Br. J.

Pharmacol. 2015, 172, 2406−2418.

(71) Strange, P. G. Use of the GTPγS ([³⁵S]GTPγS and Eu-GTPγS)

Binding Assay for Analysis of Ligand Potency and Efficacy at G

Protein-Coupled Receptors. Br. J. Pharmacol. 2010, 161, 1238−1249.

(72) Ross, R. A.; Gibson, T. M.; Stevenson, L. A.; Saha, B.; Crocker,

P.; Razdan, R. K.; Pertwee, R. G. Structural Determinants of the Partial

Agonist-Inverse Agonist Properties of 6′-Azidohex-2′-yne-Δ⁸-tetrahy-

drocannabinol at Cannabinoid Receptors. Br. J. Pharmacol. 1999, 128,

735−743.

(73) Dodd, P. R.; Hardy, J. A.; Oakley, A. E.; Edwardson, J. A.; Perry,

E. K.; Delaunoy, J. P. A Rapid Method for Preparing Synaptosomes:

Comparison, with Alternative Procedures. Brain Res. 1981, 226, 107−

118.

(56) Fichna, J.; Bawa, M.; Thakur, G. A.; Tichkule, R.; Makriyannis,

A.; McCafferty, D. M.; Sharkey, K. A.; Storr, M. Cannabinoids

Alleviate Experimentally Induced Intestinal Inflammation by Acting at

Central and Peripheral Receptors. PLoS One 2014, 9, e109115.

(57) Li, C.; Xu, W.; Vadivel, S. K.; Fan, P.; Makriyannis, A. High

Affinity Electrophilic and Photoactivatable Covalent Endocannabinoid

Probes for the CB1 Receptor. J. Med. Chem. 2005, 48, 6423−6429.

(58) Blankman, J. L.; Cravatt, B. F. Chemical Probes of

Endocannabinoid Metabolism. Pharmacol. Rev. 2013, 65, 849−871.

(59) Ahn, K. H.; Mahmoud, M. M.; Samala, S.; Lu, D.; Kendall, D. A.

Profiling Two Indole-2-carboxamides for Allosteric Modulation of the

CB1 Receptor. J. Neurochem. 2013, 124, 584−589.

(60) Cawston, E. E.; Connor, M.; Di Marzo, V.; Silvestri, R.; Glass,

M. Distinct Temporal Fingerprint for Cyclic Adenosine Mono-

phosphate (cAMP) Signaling of Indole-2-carboxamides as Allosteric

Modulators of the Cannabinoid Receptors. J. Med. Chem. 2015, 58,

5979.

(61) German, N.; Decker, A. M.; Gilmour, B. P.; Gay, E. A.; Wiley, J.

L.; Thomas, B. F.; Zhang, Y. Diarylureas as Allosteric Modulators of

the Cannabinoid CB1 Receptor: Structure−Activity Relationship

Studies on 1-(4-Chlorophenyl)-3-{3-[6-(pyrrolidin-1-yl)pyridin-2-yl]-

phenyl}urea (PSNCBAM-1). J. Med. Chem. 2014, 57, 7758−7769.

(62) Khurana, L.; Ali, H. I.; Olszewska, T.; Ahn, K. H.; Damaraju, A.;

Kendall, D. A.; Lu, D. Optimization of Chemical Functionalities of

Indole-2-carboxamides To Improve Allosteric Parameters for the

Cannabinoid Receptor 1 (CB1). J. Med. Chem. 2014, 57, 3040−3052.

(63) Mahmoud, M. M.; Ali, H. I.; Ahn, K. H.; Damaraju, A.; Samala,

S.; Pulipati, V. K.; Kolluru, S.; Kendall, D. A.; Lu, D. Structure−

Activity Relationship Study of Indole-2-carboxamides Identifies a

Potent Allosteric Modulator for the Cannabinoid Receptor 1 (CB1). J.

Med. Chem. 2013, 56, 7965−7975.

(64) Piscitelli, F.; Ligresti, A.; La Regina, G.; Coluccia, A.; Morera, L.;

Allara, M.; Novellino, E.; Di Marzo, V.; Silvestri, R. Indole-2-

carboxamides as Allosteric Modulators of the Cannabinoid CB(1)

Receptor. J. Med. Chem. 2012, 55, 5627−5631.

(65) Kumar, D.; Raj, K. K.; Bailey, M.; Alling, T.; Parish, T.; Rawat,

D. S. Antimycobacterial Activity Evaluation, Time-Kill Kinetic and 3D-

QSAR Study of C-(3-aminomethyl-cyclohexyl)-methylamine Deriva-

tives. Bioorg. Med. Chem. Lett. 2013, 23, 1365−1369.

(66) Benington, F.; Morin, R. D.; Clark, L. C., Jr. Mescaline Analogs.

V. p-Dialkylamino-β-phenethylamines and 9-(β-Aminoethyl)julolidine.

J. Org. Chem. 1956, 21, 1470−1472.

(67) Meyer, M. D.; Kruse, L. I. Ergoline Synthons: Synthesis of 3,4-

Dihydro-6-methoxybenz[cd]indol-5(1H)-one (6-methoxy-Uhle’s ke-

tone) and 3,4-Dihydrobenz[cd]indol-5(1H)-one (Uhle’s ketone) via a

Novel Decarboxylation of Indole-2-carboxylates. J. Org. Chem. 1984,

49, 3195−3199.

Q

DOI: 10.1021/acs.jmedchem.5b01303

J. Med. Chem. XXXX, XXX, XXX−XXX

Article Doi

Novel Electrophilic and Photoaffinity Covalent Probes for Mapping the Cannabinoid 1 Receptor Allosteric Site(s)

DOI: 10.1021/acs.jmedchem.5b01303

Source and publish data:

Authors:

Article abstract of DOI:10.1021/acs.jmedchem.5b01303

Full text of DOI:10.1021/acs.jmedchem.5b01303

Products guided by the article

R&D Labs maybe for 1446648-75-4

Relevant to this article

Hot Product